Public Health Mycobacteriology A Guide For The Level lll Laboratory Patricia T. Kent, B.S. George P. Kubica, Ph.D Division of Laboratory Training and Consultation Laboratory Program Office 1985 U.S. DEPARTMENT OF HEALTH AND HUMAN SERVICES Public Health Service Centers for Disease Control Atlanta, Georgia 30333 Use of trade names is for identification only and does not constitute endorsement by the Public Health Service or by the U.S. Department of Health and Human Services. Contents Introduction References Safety in the Laboratory _ A. B. C. D Specimen Collection and Transportation _ A. Laboratory Arrangement and Care — Recommended Safety Equipment and Supplies Personnel _ In Case of an Accident 1. One-Pass Air Handling System 2. Recirculating Air Handling System _ References Collection 1. Pulmonary Disease 2. Extrapulmonary Disease Transportation _ References Isolation Procedures A. B. Centrifugal Efficiency and Digestant Toxicity Digestion-Decontamination Procedures 1. N-Acetyl-L-Cysteine-Sodium Hydroxide Method Zephiran-Trisodium Phosphate (Z-TSP) Method Petroff’'s Sodium Hydroxide (NaOH) Method Oxalic Acid Method Sulfuric Acid Method s Cetylpyridinium Chloride- Sodium Chloride (CPC) Method References Culture Media 1. Advantages and Disadvantages of Egg-Base vs. Agar-Base 2. Preparation of Egg-Base Medium 3 Preparation of Agar-Base Medium 4. Preparation of Middlebrook 7H-9 Liquid Medium 5. Storage of Media Guidelines for Optimal Culture Growth 1. Preparation of Media 2. Inoculation of Media a 3 4 5 6. R PO AWN Incubation of Cultures EE Atmosphere for Incubation Temperature of Incubation Precautions eferences 15 16 16 19 20 21 21 22 23 25 29 31 31 36 36 40 42 43 44 44 46 a7 47 48 49 51 52 52 - 62 52 53 53 55 55 56 Identification Test Techniques Acid-Fast Microscopy 1. General Comments and Precautions Z2 Smear Preparation 3 Sodium Hypochlorite Method for Concentration of Mycobacteria 4. Staining Methods i i a. Basic Fuchsin Acid- Fast ‘Stains oo b. Fluorochrome Acid-Fast Stains Examination of Smears Reporting Smear Results 5 6. 7. Detecting the Source of False-Positive Smears — R eferences Introduction A. Arylsulfatase - 57 1. Centers for Disease Control Method 2. Wayne's Phenolphthalein Sulfatase Test References Carbon Sources References Catalase 1. Semiquantitative Catalase Test 2. Heat Stable Catalase Test References Growth Rate References Iron Uptake 1. Modified Method of Tison ” al. 2. Modified Method of Szabo and Vandra References Growth on MacConkey Agar Without Crystal Violet oo References Niacin Production 1. Niacin Test With Chemical Reagents 2: Niacin Test With Paper Strips References Nitrate Reduction Tests 1: Classical Method With Liquid Reagent 2. Method With Crystalline Reagents 3. Nitrate Paper-Strip Method 4. Combined Niacin - Nitrate Test References Pigment Production References Pyrazinamidase References Sodium Chloride Tolerance References Tellurite Reduction ~~ References 57 59 59 60 60 62 64 66 68 69 71 71 73 74 77 78 79 80 80 81 82 84 84 86 86 86 89 89 90 91 91 92 94 96 96 97 99 100 101 ~103 104 107 107 109 109 110 110 112 M. Thiophen-2-Carboxylic Acid Hydrazide Susceptibility Test _ — SE 112 References ~ ——— 113 N. Tween Hydrolysis re 114 References rE — 116 O. Tween Opacity 116 References SE 117 P. Urease EE } 117 1. Murphy- Hawkins Disk Method 117 2. Steadham Method: Texas Urease Test 118 3. Wayne Method ene 2 119 References _ 120 Culture Examination and Identification _ _ 121 A. Rapid Growers _ mess 129 B. Photochromogens et 133 C. Nonphotochromogens ~ Er 137 D. Scotochromogens SE 142 References SE 146 Antituberculosis Chemotherapy and Drug Susceptibility Testing 159 A. Preventive Therapy (or Chemoprophylaxis) for Tubercu- losis Infection 159 B. Chemotherapy for Disease sm 160 C. Drug Susceptibility Tests === — 162 1. Difficult to Standardize EE 162 2. Who Performs Drug Susceptibility Tests? 162 3 When to Perform Drug Susceptibility Tests 163 4. Drug Resistance oo : 163 5. Methods ee 165 6. Type of Susceptibility Test I 166 a. Direct Test ~ S— 166 b. Indirect Test —— 168 7. Preparation of Drugs and Drug-Containing Media ~169 8. Storage of Drugs _ 174 9. Disc Method E— 175 10. Reporting Test Results — 177 11. Radiometric Methods oo 181 12. Drug Assay E 181 References a — 182 Reporting Culture Results 185 Quality Control in the Mycobacteriology Laboratory 189 A. General Recommendations a 190 B. Laboratory Arrangement and Personnel 190 C. Laboratory Equipment ee 190 D. Media, Reagents, and Biochemical Tests 195 References oo 202 Appendix _ _ _ 203 Tables 5A. 5B. 6A. 6B. 7A. 7B. 8A. 8B. 9A. 9B. 10. 11. 12. 13. 14. Preparation of NALC-NaOH digestant-decontaminant solution i — Smear evaluation } EU i Numbers of indicated species of Mycobacteria recovered by State health departments of the United States Identification of clinically important Mycobacteria Rapid growers Problems in differential identification-Rapid growers Niacin positive nonchromogens Problems in differential identification-TB complex Photochromogens Problems in differential identification-Photochromogens Nonphotochromogens . Problems in differential identification-Nonphotochromogens Scotochromogens a Problems in differential identification-Scotochromogens Critical concentrations of antituberculosis drugs in Middlebrook 7H-10 medium Preparation of drug media Drug-containing discs for susceptibility tests Monitoring of equipment Quality control procedures vi Page 36 67 124 125 129 130 132 132 134 135 138 139 141 144 164 174 175 191 196 Figures Page 1. Laboratory suite Ee) _ 7 2. Removal of airborne particles Er —————— 8 3. Isolation room . _ ~ SE 9 4. Biological safety cabinets SE EI oo 10 5. Several types of safety pipettors oo 12 6. Splash proof containers _ eR oo — 13 7. Alcohol sand flask oo 13 8. Transferspades Er EE 14 9. Packaging and labeling of etiologic agents } _ 27 10. Efficiency of centrifugation - EE _ 33 11. Combined lethal effect on Mycobacteria of temperature and time of exposure to a potentially lethal digestant I in 34 12. Hypothetical examples of overall kill with different digestants ~~ I ER I 35 13. Diagram for tenfold dilutions — — 38 14. Mylar bag CO, incubator _ _—— igi 54 15. Smear area _ I rr) I 57 16. Recommended methods for examining acid-fast stained smears : _ FE 64 17. Separation of Mycobacteria on the sole basis of growth rate and pigment production 122 18. Separation by growth rate and pigment 128 19. Nonphotochromogens-separation of pathogens and saprophytes using Tween hydrolysistest ~~ 137 20. Scotochromogens-separation of pathogens and saprophytes using Tween hydrolysistest ~~ 143 21. Drugs affecting special TB populations 161 22. Drug susceptibility test medium made with drug-impregnated discs eoeee— EE 176 23. Drug susceptibility test _ 179 24. Drug susceptibility test : 180 vii Color Plates Microscopy 1. 2. 3. Branching filaments with beading and small coccoid forms Alternating clear and pink-stained bacilli that show... banding Strands of bacilli in cords Biochemical Test Reactions Arylsulfatase - CDC method Color standards for arylsulfatase test Wayne's phenolphthalein sulfatase test Semiquantitative catalase test Heat stable catalase test Iron uptake Growth on MacConkey without crystal violet Niacin chemical test Niacin paper strip test Nitrate reduction test with liquid reagents Combined niacin-nitrate test Nitrate color standards Nonphotochromogen - pigment production Photochromogen - pigment production Scotochromogen - pigment production Pyrazinamidase Tellurite reduction test Tween hydrolysis Urease - Murphy Hawkins Disk Method Urease - Steadham Method Urease - Wayne Method Colony morphology 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 35D. 36. 37. M. tuberculosis on agar and egg media M. simiae colonies on agar and egg media M. bovis colonies on agar and egg media M. asiaticum smooth colonies on egg and agar media M. kansasiicolonies (rough and smooth) on agar and egg media marinum colonies on agar and egg media . gastri colony on egg medium avium colony types on agar and egg media malmoense colonies on agar medium SXRRRRRR terrae - M. triviale colonies on agar and egg media . gordonae smooth colonies on agar and egg media . scrofulaceum smooth colonies on agar and egg media szulgai colonies (smooth and rough) on agar and egg media M. xenopi colonies on agar M. fortuitum colonies (rough and smooth) on agar and egg media M. chelonae colonies on agar and egg media viii Page 65 147 , 147 148 148 149 150 151 152 152 153 154 154 155 155 156 157 2 Introduction Introduction Over the past 15 years there has been a trend to treat tuberculosis patients in general hospitals and even in outpatient clinics, rather than in a sanatorium. This decentralization of treatment and labora- tory services led to an official statement by the American Thoracic Society (1,2) that recommended three different levels of laboratory service, the components of which were detailed in other publications (3,4). The three levels of service (I, Il, and Ill) are roughly equivalent to “Extents 2, 3, and 4,” respectively, used by the College of American Pathologists in their proficiency testing programs. The rationale behind the “‘Levels of Service” concept was that decen- tralization of tuberculosis management services might result in decreased proficiency, particularly of laboratory services, as more and more laboratories received fewer and fewer specimens. The cur- rent recognition of 54 species of mycobacteria, the need for improved isolation methods to recover these from clinical or environmental specimens, the multiplicity of in vitro tests required to speciate the isolates, and the requests for susceptibility tests against a broad range of chemotherapeutic agents all demand a level of laboratory exper- tise that cannot easily be learned in today’s atmosphere of decentral- ized management of tuberculosis. The training programs in mycobacteriology conducted by the Cen- ters for Disease Control (CDC) since 1975 were designed with the “Levels” concept in mind. Brief summaries of the functions performed at each level of service are shown below. The relegation of a labora- tory to a given level of laboratory service is through self-evaluation. Directors may find that some tests performed by their personnel are done so infrequently that results are not credible. They may then decide to refer these tests to another institution or to upgrade the work of their personnel through training and proficiency testing. Before such decisions are made, the director should consider the following questions: What is the frequency of a requested service? Does the work volume in mycobacteriology justify the time, effort, and expense allocated to that service? Am | confident of the results we provide the clinician? Do our facilities have the required space, equipment, and airflow to ensure the safe handling of specimens? Remember, the clinician relies upon laboratory results to establish the diagnosis of mycobacterial disease and often to designate cura- tive therapy for the patient. It is important that these results are accu- Level Type of Facility* Type of Service | Physician's Office Collect good specimens. Ship to Outpatient Clinic Levels Il or Ill for culture. Small Hospitals Perform microscopic examination. Il Treatment Center Collect specimens. Selected Laboratories Perform microscopic examination. (City, County, State, Isolate organisms in culture. or Private) Identify Mycobacterium tuberculosis. Perform susceptibility tests. ** Refer “other” mycobacterial isolates to Level lll for identification. I} Reference Laboratory Perform all procedures of Level Il. (State, Federal, Private) Identify all mycobacteria. Perform susceptibility tests. ** * Examples of Type of Facility are suggestive, not directive. ** Laboratory personnel should not perform susceptibility tests unless they (a) can identify the organism they are testing and (b) perform a sufficient number of such tests to be cognizant of the many problems associated with the procedure. rate and dependable, so the need for an honest, self-appraisal of laboratory work performance is obvious. To assist laboratory personnel who ultimately assign themselves to one of the three levels of expertise, CDC staff have prepared three separate training courses, one for each level of laboratory service. The “Workshop in Recognition and Quantitation of Tubercle Bacilli in the Level | Laboratory” is designed for those laboratories that collect specimens and may be asked to perform quantitative micro- scopic examination of stained smears. A separate manual on Acid- Fast Microscopy is available (5). It is suggested that proficiency in smear examination may be maintained by laboratories that examine 10 to 15 smears per week, provided adequate controls are used. If the training materials for this course are not available on loan from your State health department, they may be borrowed from the Division of Laboratory Training and Consultation (LTC), Laboratory Program Office (LPO), CDC. The field course “Isolation and Identification of Mycobacterium tuberculosis” may be requested through the State health department by those laboratories that function at Level Il and that routinely pro- cess and culture 20 clinical specimens per week to ensure proficiency. Although this course is commonly presented by CDC staff, LTC per- sonnel will work with those States that prefer to present the course themselves. A separate manual addresses the activities of the Level Il laboratory (6). This manual was prepared as a guide for the Level lll course con- ducted at CDC, and for those laboratories involved in reference mycobacteriology work. Although all 54 species of mycobacteria rec- ognized in 1983 are mentioned, attention is directed only to about 20 of those most commonly encountered in the reference laboratories. Detailed directions are presented for performing procedures that have proved successful at CDC for isolation, differential identification, and drug susceptibility testing of mycobacteria. On occasion, alternative procedures that have proved satisfactory in other institutions are also described; readers may select those that best meet their needs. Selected references at the end of most sections provide additional detail or critical evaluation of procedures described in this text. It is impossible to acknowledge all who have provided advice, encouragement, and assistance in the preparation of this manual. We hope you forgive our unintentional omissions. Especial thanks go to Dr. Robert C. Good, Mrs. Vella A. Silcox, Mr. Ronald W. Smithwick, and Dr. James O. Kilburn of the Mycobacteriology Branch, and Dr. Dixie E. Snider, Jr., of the Division of Tuberculosis Control, CDC, for review, criticism, and contributions to the text; to Dr. Nancy G. Warren, Consolidated Laboratory Services, Department of General Services, Commonwealth of Virginia, for helpful suggestions and criti- cism borne both of work experience and an attempt to put to service the things described in this and similar manuals; and to Mr. A. Ray- mond Simons, LTC, LPO, for the many new color plates that have been added to this 1984 edition of Public Health Mycobacteriology - A Guide for the Level lll Laboratory.” A special note of gratitude is due Mrs. Miriam Dugger for her patience and perseverance in setting our ideas into manuscript form. REFERENCES 1. American Thoracic Society. Quality of laboratory services for mycobacterial diseases: official statement of the American Thoracic Society. Am Rev Respir Dis 1974;110:376. 2. American Thoracic Society. Levels of laboratory services for mycobacterial diseases: Official Statement of the American Thoracic Society. Am Rev Respir Dis 1983;128: 213. 3. Hawkins JE, Good RC, Kubica GP, Gangadharam PR, Gruft HM, Stottmeier KD, Sommers HM, Wayne LG. The levels of service concept in mycobacteriology. Posi- tion paper adopted by the American Thoracic Society. ATS News, Summer 1983: 19-25. (Copies of this and reference 2 available from State and local Lung Associa- tions). 4. Kubica GP, Gross WH, Hawkins JE, Sommers HM, Vestal AL, Wayne LG. Laboratory services for mycobacterial diseases. Am Rev Respir Dis 1975; 112:773-87. 5. Smithwick RW. Laboratory manual for acid-fast microscopy, 2nd ed. Atlanta: Center for Disease Control, PHS, HEW, 1976. 6. Strong BE, Kubica GP. Isolation and identification of Mycobacterium tuberculosis: a guide for the level Il laboratory. Atlanta: Center for Disease Control, PHS, HEW, 1981. Available from U.S. Dept. of Commerce, National Technical Information Service, 5285 Port Royal Road, Springfield, VA 22161 (Publication No. PB82-164625; Cost $15.50). Safety in the Laboratory Safety in the Laboratory Work-related infections are a recognized hazard for personnel employed in laboratories where infectious disease agents are han- dled (13). Studies have shown the risk of tuberculous infection to be three to five times greater for the mycobacteriology laboratory worker than for other personnel (secretaries, maintenance workers, etc.) in the same institution (8,14). Mycobacterium tuberculosis produces a chronic disease curable only after 6 to 18 months of drug therapy. Other potentially pathogenic mycobacteria commonly exhibit a high level of resistance to most antituberculosis drugs that makes success- ful treatment unpredictable. Although there is little or no evidence of laboratory-associated infections with these other mycobacteria, we must assume that they occur. To believe otherwise would suggest that M. tuberculosis is uniquely capable of infecting laboratorians by inhalation, unintentional injection (as with contaminated, broken glassware), or any other route. Because these other mycobacteria collectively account for about half of the acid-fast cultures recovered in the laboratories of the United States, we must assume that they, just as M. tuberculosis, have the same chance of infecting humans via the same routes by which tubercle bacilli gain access to the body. With these facts in mind, it is obvious that safety in the mycobacteri- ology laboratory must receive major emphasis, and the reader is urged to apply the biosafety principles set forth in the CDC-NIH guide- lines (2). Safety in the laboratory must start at the administrative level. If the organization is large enough, there should be an Office of Biosafety or at least a Biosafety Officer who has the demonstrated administra- tive support for a safety program. There are, of course, many facets of a laboratory safety program, some of which will be addressed here only peripherally. Our primary safety objective in this manual is to suggest ways to'minimize laboratory-related infection with mycobac- teria. It is an administrative responsibility to ensure that the employee is (a) monitored regularly by medical personnel; (b) trained properly in safe laboratory procedures; (c) informed of especially dangerous tech- niques and procedures that might require special care; (d) prepared for prompt and correct action following an unexpected accident; and (e) provided with adequate safety equipment. Each of these responsi- bilities is discussed in this chapter. The laboratory worker must be responsible for using the appropriate safety equipment, following established laboratory policy, and accepting responsibility for correct work performance to assure the safety of all. Most infections in the mycobacteriology laboratory can be attrib- uted to the unrecognized production of potentially infectious aero- sols containing acid-fast bacilli (13). Although some aerosolized drop- lets are large (>5 um) and settle rapidly to contaminate skin, clothing, and countertops, the most dangerous aerosols are those that pro- duce droplet nuclei, tiny dry particles <5 um in size (that may contain 1 or 2 viable mycobacteria) that remain suspended almost indefi- nitely in air unless they are removed by controlled airflow or ventila- tion (5,6,16). If these tiny, infectious droplet nuclei are not controlled or eliminated, they are capable of entering a pulmonary alveolus and establishing the primary site of infection. Studies have shown that the following laboratory manipulations (9,17) yield aerosols composed primarily of droplet nuclei: e Pouring of liquid cultures or supernatant fluids e Using fixed volume automatic pipettors e Mixing a fluid culture with a pipette e Using a high-speed blender e Dropping tubes or flasks of broth cultures e Breaking tubes during centrifugation e Using paint conditioning machines, as for processing sputum e Letting drops of microbial suspension fall from a pipette onto a hard work surface Although laboratory-associated infections are usually due to the inhalation into the lung of droplet nuclei containing mycobacteria, there are other less common routes of infection: self-inoculation with needle and syringe; puncture of skin with broken, contaminated glassware; and infection through uncovered cuts or abrasions. Because of these extra-pulmonary routes of infection, Good House- keeping Is Mandatory! All surfaces and equipment within the isola- tion room (including the biological safety cabinet) should be regarded as potentially infectious and should be cleaned regularly by appropri- ate means; e.g., disinfectant swabbing, autoclaving, ultraviolet light. The following descriptions and recommendations suggest ways (a) to minimize aerosol production during routine laboratory manipula- tions and (b) to eliminate quickly those aerosols that do form. A. Laboratory Arrangement and Care Ideally, all mycobacteriology work should be performed in a self- contained suite that houses all supplies and equipment needed to process specimens, from time of receipt to final report (18). Only laboratories that have a class | or class Il biological safety cabinet (BSC) should perform mycobacterial examinations. The suite should be constructed with a one-pass (nonrecirculating) ventilation system to maintain airflow in one direction only, from a clean to a less clean area, as indicated by the arrows in figure 1. The highest air pressure (+ +) should exist in the corridor, while the general laboratory is under more negative (—) pressure. In this way, the air always flows into the laboratory from the corridor and not in the opposite direction. If there is a sterile room (e.g., for media preparation) in the laboratory suite, it should be under more positive pressure (+ —) than the general laboratory area (—) to keep airborne contaminants from moving into the sterile room. The isolation room should be under the most negative pressure (——) with its exhaust air moving through the thimble unit or through high efficiency particu- late air (HEPA) filters in the BSC before the exhaust air ducts (———) carry it to the outside. Be aware of the total number of room air changes per hour and the airflow pattern in your laboratory. Know if the laboratory has a one-pass or a recirculated air handling system (information may be obtained from maintenance personnel). Six to EXHAUST DUCT (——-) pcr eee ] F——-=."FE———] Biological rrr 9 Safety Cabinet ISOLATION ROOM (——) z a oc oc i 8 Auto- SS clave I 2 RN ~~ Bench or Space for Movable Equipment or Furniture Sink with — Foot-Operated Valves GENERAL LABORATORY (—) (++) STERILE ROOM (+—) Bench FIGURE 1. Laboratory suite Air pressure indications: ——— (least) to ++ (greatest) twelve room air changes per hour are very good and provide removal of up to 99% or more of airborne particulates within 30 to 45 minutes (figure 2); published figures of 20 changes per hour (1), although effective, would probably create greater problems of air turbulence within the BSC and the laboratory. Place short strips of tissue paper on air duct grills located in doors and walls, and at the edge of the BSC opening, to monitor constantly the direction of airflow. Work should never be performed when the suite is not under negative pressure (i.e., reversal of airflow direction or no airflow at all). The air from the suite should be vented to the outside through a dedicated exhaust duct or a nonrecirculating build- ing exhaust system. Exhaust air should be discharged high up on the building where there is little possibility of it being reintroduced into the building (i.e., do not place air intake and exhaust in proximity to one another) (2). The “Isolation Room" (figure 3), containing a BSC, should be a separate room within the suite. All procedures that may generate aerosols should be performed within the BSC (e.g., opening Air Changes | Minimum time for reduction of airborne Per Hour particles by indicated percentage 99% 99.9% 4 69 104 — 46 69 —e= 10 28 1 W -==-= 15 18 28 \ \ Formula for 46 6.9 \ \ \ time (hours) ——————— | ———————— 90+ , “ needed to reduce Number of air Number of air \ airborne particles] changes per hour | changes per hour \ \ vv S by indicated vo \ percentage : \ = 99 —| 8 © a E § = 99.9 Q 3 ° oc @ > & § 99.99 © a Time in Minutes FIGURE 2. Removal of airborne particles and processing clinical specimens, preparing smears, inoculating or transferring cultures, and performing in vitro tests). The exhaust air from the BSC must be discharged through HEPA filters that have been tested and certified to remove 99.97% of particulates (0.3 wm or larger). Fe ee] ) —— — BIOLOGICAL SAFETY CABINET CHAIR AUXILIARY WORK TABLE ~ ‘0 © © WATER BATHS ISOLATION ROOM DISCARD HAMPER TABLE -_es ees ees] FEET FIGURE 3. Isolation room B. Recommended Safety Equipment and Supplies General maintenance or repair in the mycobacteriology suite should be performed under careful supervision. Isolation room equipment should not be serviced, checked, or cleaned unless a trained person is present to ensure that adequate safety precautions are observed. When major servicing is needed, the entire room and the BSC should be decontaminated (see ““D. IN CASE OF AN ACCIDENT,” which follows). If only a single piece of equipment requires servicing, it may be decontaminated with a tuberculocidal disinfectant and removed from the isolation room. Biological Safety Cabinet The single, most important piece of laboratory equipment needed in a mycobacteriology laboratory is a well-maintained, properly func- tioning biological safety cabinet (BSC). Descriptive manuals of the construction, use, maintenance, and decontamination of biological safety cabinets are available (2, 10, 11, 12, 18,). These manuals, together with the manufacturer's brochure, should be available to laboratory staff and maintenance personnel. Processing raw or “untreated’’ clinical specimens or transferring viable cultures should not be permitted in a laboratory that does not have a BSC. Two types of cabinets are in use and are satisfactory (see figure 4). One is a class | negative-pressure biological safety cabinet (NPBSC) that draws a minimum of 75 linear feet of air per minute across the frontal opening and exhausts 100% of the air to the outside. The other is a class II vertical laminar flow biological safety cabinet (LFBSC) that blows HEPA filtered air over the work area. The models A, B1 and B3 that recirculate 70% and 30% of the air are suitable for use in the myco- bacteriology laboratory. Model B2 is designed for “‘total (100%) exhaust’ and is not required in the mycobacteriology laboratory (12). It is strongly recommended that a BSC should not be installed in a mycobacteriology laboratory unless it is vented into a nonrecirculating exhaust system or directly to the outside. The airflow through the BSC is adjusted by the manufacturer to provide at least 75 linear feet per minute (Ifpm), and should be tested and recertified at least yearly by trained personnel. Periodic (e.g., quarterly or more often under dusty conditions) checks on the airflow may be made with an anemometer. If airflow is appreciably dimin- ished, this may indicate that filters have become clogged. Either decon- / % J : 0 Class | In In 1 n Type - A B1 B2 B3 Inflow (Ifpm) 75 75 100 100 100 % Recirculation 0 70 30 0 70 Downflow (Ifpm) 0 75 50 80 50 Exhaust (cfm/ft) 50 a5 65 65 65 Use: Biohazards + + + + + Toxic, Volatiles + —- + + + Protection u u/p U/P u/p u/P (User/Product) FIGURE 4. Airflow characteristics of Class | (negative pressure) and Class Il (vertical laminar flow)* biological safety cabinets * National Sanitation Foundation Standard 49, Revised May 1983 10 taminate the BSC with formaldehyde or purchase this service con- tractually before removing the filters for replacement (2,11). The speed of the exhaust fan on some BSCs may be increased to maintain minimal face velocity of 75 Ifpm without the need for a complete decontamination. Ultraviolet Lights Ultraviolet (UV) lights emitting rays primarily of wavelength 254 nm are effective for decontaminating work surfaces (e.g., after work is completed in the BSC) and killing airborne microorganisms (4,15). Bulbs may be mounted inside the work area of the BSC and in shielded wall or ceiling units to irradiate the upper air of the isolation room (15, 16). If wall and ceiling units are properly shielded so that radia- tion at “work level” does not exceed 0.1 wu watt/cm?, there is little or no danger to human health, and these fixtures may be allowed to burn continuously (14a, 16). Air movement through the BSC, however, is so rapid that internally mounted UV lights are of little value, except possibly for postwork decontamination of surfaces. Care should be taken when working with such lights inside the BSC. Do not work in the BSC with UV lights on or look directly at the burning bulbs because direct or reflected UV light can cause severe burns to the eyes and to other exposed parts of the body. The UV light left burning inside the BSC during work manipulations can kill mycobacteria that are being cultured or transferred. Turn UV lights on for a minimum of 1 hour after work is completed. Continuous operation when the cabinet is not in use costs very little and may be more convenient. The ultravio- let light has very little penetrating power and is easily blocked by such substances as dust and grease. Clean bulbs frequently with alcohol-soaked gauze and check germicidal output at least every 3 months. Replace the bulbs when the initial output is decreased by 70%. New UV monitor cards (Vanguard International, 1111-A Green Grove Road, Neptune, NJ 07753) are available that have spots that turn red after accumulated doses of 16,000 p watt-sec/cm? and 150,000 uw watt-sec/cm? of UV energy have accumulated at the sites. These may prove to be useful and easy methods to monitor the efficiency of UV lamps. Protective Clothing Since no BSC is 100% effective and both physical and mechanical failures do occur, protective clothing (especially the face mask) pro- vides an additional measure of personnel protection. Wear face masks designed to filter >90% of particles ranging from 0.5 um to 1.0 pm. To optimize filtration efficiency of masks (and increase personnel protec- tion), use metal nose tabs and mask ties to ensure a tight fit on the face. 11 Hospital surgical gowns protect the skin and clothing from large droplets greater than 5 wm that may accidentally be splattered. Rubber gloves guard against infection through cuts or abrasions on the hands. Autoclavable or disposable shoe covers and caps minimize the chance of transporting infectious agents from the isolation room to other parts of the laboratory or even to your residence. All protective clothing should be placed into covered containers or laundry bags and autoclaved before being washed or discarded. Auto- clave indicators (e.g., spore strips or homemade test systems) should be placed close to the center of clothing bags before autoclaving. Centrifuges If centrifuge tubes leak, crack, or shatter during centrifugation, an invisible cloud of potentially infectious droplet nuclei may be emitted and disseminated about the room. Therefore, all materials suspected of being potentially pathogenic should be centrifuged in commer- cially available aerosol-free carriers. Because tubes may break or leak during centrifugation, remove tubes from safety carriers only inside the BSC. An inexpensive centrifuge air-filtration system (7) provides an alternative to the sealed, aerosol-free carriers. See further discus- sion on “Centrifugal Efficiency and Digestant Toxicity’ in section on “Isolation Procedures.” Safety Pipetting Never mouth pipette. Not only is there a danger of aspirating infec- tious material, but an aerosol is created when the fluid is alternately sucked and expelled through the pipette. With the nose directly over the open container of infectious material, the aerosol has direct access to the lung (9). Safety pipettors should be used and are commercially available. Some examples are shown in figure 5. FIGURE 5. Several types of safety pipettors: (a) pipette filler, (b) Caulfield, (c) rubber tubing pipettor (homemade), (d) rubber bulb. 12 Autoclave Ideally, the autoclave should be inside the mycobacteriology suite to prevent contaminated material from being discarded or washed before decontamination. Use spore strips or some other biological monitor or physically check the system to verify that each load has been autoclaved. Splash-Proof Container After the digestion-decontamination procedure, pour the superna- tant fluid from sedimented specimens into a splash-proof container to minimize both aerosol production and contamination of the work surface of the BSC. Figure 6 illustrates two such containers. One-hole er stopper L Soldered joint 1" below 1" into side arm container Cotton plug ——— Disinfectant ————— Metal container Glass flask FIGURE 6. Splash-proof containers Alcohol-Sand Flask Before sterilizing in a flame, clean large clumps of bacilli and organic debris from wire inoculating loops or spades by stabbing them several times into a 250 ml, screwcap flask containing 95% alcohol and washed sea sand. The abrasive action of the sand removes most of the debris from the wire, and the alcohol causes rapid incineration of residual debris when the loop or spade is flamed in a Bunsen burner. See figure 7. Alternatively, single-use dis- posable loops also are available. Alcohol 1” \\ bore sand sand FIGURE 7. Alcohol sand flask 13 Culture Transfer Spade A spade is useful to transfer solid growth from one medium to another or to emulsify such growth into a liquid to prepare the heavy suspensions needed for some in vitro tests. Flatten one end of an 18-gauge nichrome wire to make an area 5to 10 mm long, 1to 2 mm wide, and 0.3 to 0.5 mm thick. Smooth the edges with a file or abra- sive stone. See figure 8. Use this with the sand flask above. An alternative to the metal transfer spade is the wooden applicator stick, 6 to 8 inches long (select appropriate size for tube length). Cut the end of the stick at a 30° to 45° angle, package in cannisters or tubes (as for pipettes) and sterilize by autoclaving. Such sticks could be used once and discarded. Because the sticks require no steriliza- tion in a Bunsen burner (as do nichrome spades), they are especially useful in LFBSC’'s where the use of heat-generating equipment is discouraged (because of heat buildup associated with recirculated air in these BSCs). Wooden Nichrome Wire Applicator Stick . : \ Front Side Cutat View View OR 30-45 Angle \ FIGURE 8. Transfer spades Disinfectants The temporal killing action of disinfectants depends on the concen- tration used, the duration of contact, the population of organisms to be killed, and the presense of organic debris. Most quaternary ammo- nium compounds are relatively ineffective against mycobacteria although they may be bacteriostatic. Some newly formulated quarter- nary ammoniums are reported to be mycobactericidal. The use of a 5% phenol solution is no longer recommended because of its docu- mented toxicity, but other phenolic derivatives are available, among them Amphyl (Lehn & Fink, Toledo, Ohio), Osyl (National Laboratories, Toledo, Ohio), Staphene and Vesphene ( Vestal Laboratories, St. Louis, 14 MO). Other effective agents include sodium hypochlorite at a concen- tration of 0.1% to 0.5% (e.g., 1/50 dilution of most household bleaches) and 3% to 8% formaldehyde. Although extremely effective, the latter two have limited usefulness because of a strong oxidizing action and an irritating, toxic odor, respectively. When bare hands become contaminated, a rinse with 70% isopropyl alcohol followed by thor- ough washing with soap and water is effective. With so many disinfec- tants available, it is important to consult the product brochures to make certain the disinfectant is bactericidal for mycobacteria. When handling infectious material, it is best to work over a disinfectant- soaked gauze pad, both to minimize spatter and aerosol formation if microbial inocula are dropped or spilled, and to decontaminate the spill. Some of the substituted phenolic disinfectants work best for this. C. Personnel Personnel should be selected with care; they should be physically and mentally capable. All new personnel should receive (a) a tubercu- lin skin test, (b) a chest X-ray, and (c) training in safety techniques and procedures for the mycobacteriology laboratory. Because a single, uniform surveillance program for laboratory personnel is difficult to establish, guidelines are offered for two types of laboratory situations: the “‘safe’’ and the “unknown.” In the “‘safe’’ laboratory: « personnel have been trained in correct laboratory procedures « equipment (especially the BSC) is functioning properly and monitored regularly « routine skin testing has revealed no tuberculin converters among the staff Surveillance in the “‘safe’’ laboratory would consist of (a) annual skin tests only for tuberculin-negative personnel; (b) X-rays only if an individual exhibits symptoms or has converted to tuberculin posi- tive (1a, 3). If there is a frequent turnover of personnel in the labora- tory (e.g., because of rotation through different laboratory services), skin tests should be done every 3 to 6 months, rather than annually. If possible, the rotation system in the mycobacteriology laboratory should be discouraged (except as a training aid) because expertise cannot be maintained and personnel are rarely able to follow a speci- men from receipt to final report. In the “unknown” laboratory, any one or all of the following may exist: « personnel have not been trained in correct laboratory proce- dures e equipment is not monitored for proper performance « little is known about the tuberculin conversion rate among the staff 15 Recommended surveillance for the ““unknown’’ laboratory would be to (a) retest tuberculin-negative staff at 4-month intervals until it is evident that all equipment is functioning properly and that staff mem- bers are adhering to safety procedures (this would be demonstrated by the lack of conversions; arbitrarily, 2 to 3 person-years of tubercu- lin negativity among laboratory workers should be evidence that good safety procedures are being practiced); (b) X-ray personnel only if they exhibit symptoms or if they convert to tuberculin positive; (c) monitor safety equipment and laboratory airflow to ensure optimal performance; and (d) discuss laboratory safety precautions with all personnel. IF AN EMPLOYEE'S SKIN TEST CONVERTS TO POSITIVE: Refer the employee to a physician for evaluation. Check the laboratory habits of the employee. Reevaluate laboratory safety equipment and procedures. Remember, even the best safety equipment is no substi- tute for meticulous technique on the part of a careful, safety-conscious bacteriologist. D. In Case of an Accident Laboratory safety doesn’t just happen. It is the result of (a) recogniz- ing that accidents can and will occur; (b) discussing ways to minimize and prevent accidents; (c) formulating a plan of action so the poten- tially harmful effects of an accident may be neutralized as rapidly and effectively as possible. Laboratory personnel should be encouraged to think about things that “could go wrong’ with each laboratory procedure they use, and to suggest ways to minimize or eliminate them. Everyone hopes (some even boast) that a laboratory accident will never happen, but the best defense against such an eventuality is a well-thought-out plan to neutralize any accident that may occur. No accident should be considered insignificant, but a great deal of per- sonal judgment is involved in the assessment of the seriousness of each accident and how it will be “neutralized.” The final decision is markedly affected by the amount of aerosol generated and the type of airhandling system in the facility. Some illustrative examples follow. 1. One-Pass Air Handling System The one-pass (nonrecirculated) air handling system provides 100% fresh air to the laboratory area and passes the potentially contami- nated air from the BSC through HEPA filters before exhausting it to the outside. Much of the room air (outside the BSC) is exhausted through a thimble exhaust duct without filtration. The one-pass air 16 system greatly minimizes the chances for infection because laboratory- generated aerosols are not recirculated in the building or in the laboratory. Most newly constructed public health laboratories are built with a one-pass (nonrecirculated) air system. Accidents that occur in such laboratories may be divided into two types: those that gener- ate minimal aerosol and those that produce a large quantity of poten- tially infectious aerosol. A minimal aerosol might be created by break- ing a single culture tube of solid medium, dropping a plastic petri dish, or spilling the contents of a sputum specimen. In such cases, the solid medium and the thick mucoid nature of the sputum specimen greatly limit large numbers of bacilli from being aerosolized. When such a ‘minor’ accident occurs, the plan of action should be (a) cover the spill immediately to prevent further aerosolization (use available toweling or even a laboratory coat); (b) soak the covering cloth with disinfectant to wet the area; (c) leave the room for at least 2 hours to permit the air handling system to evacuate most of the aerosol (see figure 2); (d) wear protective clothing (gown, mask, gloves) to reenter the room and clean up the spill; (e) place all clean-up material (broken tubes, plates, clothes) in appropriate containers and autoclave; (f) mop the floors and countertops with disinfectant. Note: to be prepared for such an accident have a supply of large cloths, and a wide-mouthed container of disinfectant (to facilitate rapid pouring) readily accessible in or near areas where accidents are most likely to occur. A large quantity of potentially infectious aerosol may be generated by breaking a flask or tube of liquid culture containing a high concen- tration of bacilli in suspension (e.g., 10% per ml). Take the following action: (1) Evacuate the room immediately. The danger from the poten- tially infectious aerosol is greater than any need to cover the spill. (2) Leave the BSC operating and do not reenter the room for at least 4 hours. This will provide considerable dilution of the infectious droplet nuclei (figure 2). Moreover, evacuation of the air through the HEPA filter system of the BSC should reduce the likelihood of people outside the building becom- ing infected. (3) If itis possible and feasible in your laboratory, decontami- nate the room(s) after the 4-hour waiting period by using formaldehyde gas. (This cannot be done in rooms with suspended ceilings, porous walls, or recirculated air; see page 19 for ways to handle such rooms). Even in well- sealed rooms, the possibility of “formaldehyde leaks” and the potential toxicity to personnel dictates that the follow- ing procedure be done after normal work hours. Because 17 36% to 40% formaldehyde (formalin) is readily available in most laboratories, this procedure will be outlined: (a) Seal all air intake and exhaust grills in the room. This may be as simple as taping large plastic garbage bags over the grills, or as sophisticated as having a flanged frame over each grill which will accept a metal or solid plastic sheet that can be taped in place to ensure tightness. Also tape around door frames or other open- ings through which the formaldehyde vapor may leak. (b) Use an electric hot plate to boil off 1 ml of formalin per cubic foot of room space. Example: A room 10x12x10 feet would have 1200 cu. ft. of space and would be treated with 1200 ml of formalin. Caution: DO NOT OVERDOSE THE ROOM WITH FOR- MALDEHYDE, BECAUSE EXPLOSIVE CONCENTRA- TIONS (i.e., >8%) CAN BE ATTAINED. THE AMOUNTS LISTED HERE ARE WELL WITHIN THE SAFETY RANGE. (c) Raise the relative humidity of the room to about 70% to ensure optimal effect of the formaldehyde. Most chemistry or physics handbooks have tables of the amount of water that can be held in a given volume of air at full saturation; you need only 70% of this amount. Calculate the quantity needed and boil it off on an electric hot plate. Example: In our 1200 cu. ft. room, the amount of water needed to fully saturate the room at 70°F (21°C) is 18.45 gm (or ml) per cubic meter (or per 35.3 cu. ft.). By simple mathematics, 1200 cu. ft. divided by 35.3 cu. ft. in 1 cubic meter x 18.45 gm/cubic meter x 0.7 (for 70% humidity) equals 439 gm (or ml) of water. If 500 ml of water is boiled off in the room at the same time (or just before) the formaldehyde is vaporized, the desired humidity should be attained. (d) Allow the formaldehyde vapor to stay in the room at least four hours or, preferably, overnight. Put on a gas mask* to enter the room and remove the covers from air intake and exhaust grills. Allow the room to air until no more formaldehyde is detectable, then mop all residue from the floors, walls, and counters. If a white, powdery residue is obvious, this may be re- moved by wiping with a 10% ammonium hydroxide solution (use gloves). * MSA Chin Type Gas Mask with Ultravue Facepiece available from Mine Safety Appliances Co., 400 Penn Center Blvd., Pittsburgh, PA 15235. Should be used with Chin Type Canister, Type GMR. 18 2. Recirculating Air Handling System Most buildings constructed in the 1960's or earlier had a recirculat- ing air system that drew in 20% fresh air to mix with 80% of pre- viously circulated air. Although such a system is less expensive to heat and cool, it is not desirable for laboratories that deal with poten- tially hazardous chemical or biological agents. If infectious aerosols are produced in the laboratory, they may be circulated throughout the building, infecting people distant from the site of the aerosol- generating accident. Because the air is recirculated, the formalde- hyde gas decontamination procedure is not feasible; nor should it be used with the one-pass air system if laboratory areas have suspended ceilings or porous (cinder block) walls. In these situations, when room decontamination is necessary, the use of the fogging machine* is recommended. A fogging machine rapidly saturates the air, causing the dangerous droplet nuclei to settle. If an appropriate disinfectant is used in the fogger, the potentially infectious droplet nuclei are also decontaminated as they settle to the floors and countertops.* The appropriate plan of action following an aerosol-generating acci- dent in a laboratory with a recirculating air handling system would be: (1) Evacuate the area immediately, except for the person who caused the accident (2) Shut off the air handling system if possible (maintenance personnel should be alerted to this possibility) (3) Seal the exhaust and intake air ducts as quickly as possible to prevent pressure from building in the room (use plastic or metal sheets and seal with tape) (4) Charge the fogging machine with disinfectant (kept pre- pared in approximate volume), turn on the machine, exit the room, and tape the door shut (5) Let the fogger dispense the entire volume of disinfectant, permit the fog to settle, and leave the room undisturbed for at least an hour (6) Put on protective clothing before reentering the room (7) Complete the cleanup by mopping the floor and counters with disinfectant soaked cloth This is a messy procedure, but rapid and effective in buildings with recirculated air, suspended ceilings, or porous walls. * Fogmaster (Germfree Laboratories, Inc., Miami, FL) and Oxford Jet Fogger (Oxford Chemicals, Inc., Chamblee, GA). 19 References 1. 1a. 10. 12. 13. 14. 14a. 15. 16. 17. 18. American Thoracic Society, Official Statement. Diagnostic standards and classifica- tion of tuberculosis and other mycobacterial diseases (14th ed.) New York: Ameri- can Thoracic Society, 1981. Bureau of Radiological Health. The national conference on referral criteria for x-ray examinations. Washington, DC: Food and Drug Administration, PHS, HEW, 1979 (HEW publication no. (FDA) 79-8083). . Centers for Disease Control and National Institutes of Health. Biosafety in microbio- logical and biomedical laboratories. Atlanta: Centers for Disease Control, PHS, HHS, 1984. (Available from Superintendent of Documents, U.S. Government Print- ing Office, Washington, DC 20402, as HHS Publication No (CDC) 84-8395.) . Cope R, Hartstein Al. The annual chest roentgenogram for control of tuberculosis in hospital employees: recent changes and their implications. Am Rev Respir Dis 1982;125:106-7. . David HL. Response of mycobacteria to ultraviolet light radiation. Am Rev Respir Dis 1973;108:1175-85. . Foord N, Lidwell OM. a. Airborne infections in a fully air-conditioned hospital. |. Air transfer between rooms. J Hyg 1975; 75:15-30. . Foord N, Lidwell OM. b. Airborne infections in a fully air-conditioned hospital. Il. Transfer of airborne particles between rooms resulting from the movement of air from one room to another. J Hyg 1975; 75:31-44. . Hall CV. A biological safety centrifuge. Health Lab Sci 1975; 12:104-6. . Harrington JM, Shannon HS. Incidence of tuberculosis, hepatitis, brucellosis, and shigellosis in British medical laboratory workers. Br Med J 1976;1:759-62. . Kenny MT, Sabel FL. Particle size distribution of Serratia marcescens aerosols created during common laboratory procedures and simulated laboratory accidents. Appl Microbiol 1968;16:1146-50. National Institutes of Health. Certification of class Il (laminar flow) biological safety cabinets. Bethesda: National Cancer Institute, NIH, PHS, HHS, 1975. . National Institutes of Health. Formaldehyde decontamination of laminar flow bio- logical safety cabinets. Bethesda: National Cancer Institute, NIH, PHS, HHS, 1984. National Sanitation Foundation. Standard no. 49 for class Il (laminar flow) biohazard cabinetry. Ann Arbor, MI: National Sanitation Foundation, 1976 (revised 1983). Pike RM. Laboratory-associated infections: summary and analysis of 3921 cases. Health Lab Sci 1976;13:105-14. Reid DP. Incidence of tuberculosis among workers in medical laboratories. Br Med J 1957;2:10-14. Riley RL. Airborne infection. Am J Med 1974,57:466-75. Riley RL, Knight M, Middlebrook G. Ultraviolet susceptibility of BCG and virulent tubercle bacilli. Am Rev Respir Dis 1976;113:413-18. Riley RL O'Grady F. Airborne infections, transmission and control. New York: McMillan Company, 1961. Stern EL, Johnson JW, Vesley D, Halbert MM, Williams LE, Blume P. Aerosol production associated with clinical laboratory procedures. Am J Clin Pathol 1974;62:591-600. Strong BE, Kubica GP . Isolation and identification of Mycobacterium tuberculosis: a guide for the level Il laboratory. Atlanta: Centers for Disease Control, PHS, HHS, 1981. 20 Specimen Collection and Transportation Specimen Collection and Transportation The presence of acid-fast bacilli in a clinical specimen may be con- firmed either by microscopy or by cultural examination. Since mycobacteria cannot be specifically identified by smear examination alone and because there are many species known to infest or infect man, the definitive diagnosis of a mycobacteriosis (other than leprosy) can be made only if the infecting Mycobacterium is isolated from the submitted clinical specimen. The efficiency of any laboratory proce- dure used to culture mycobacteria from clinical specimens depends on the manner in which the specimen is obtained and handled. Therefore, specimens should be collected with the utmost care and promptly transported to the laboratory. A. Collection For optimal results, obtain clinical specimens under the following conditions: « Collect specimens before chemotherapy is started; even a few days of drug therapy may kill or inhibit sufficient numbers of acid-fast bacilli to leave confirmation of disease in doubt. e Collect specimens in clean, sterile, one-use, plastic, dispos- able containers. If this is not possible, use glass containers that have been washed with dichromate sulfuric acid and sterilized. e Collect a series of three to six single, early morning sputum specimens on successive days. The single specimen is easier to handle in the laboratory and less likely to be contaminated, especially if sent by mail (4,7,9,13,15,20). « Transport specimens to the laboratory as soon as possible. Refrigerate specimens if delivery is delayed (7). « Seal and package specimen containers carefully to avoid leak- age or breakage in transit (17). The staff responsible for collecting and transporting specimens should be provided an instruction sheet that outlines the proper procedures. The attending medical personnel should be notified of any deviation from the acceptable procedure that might result in a less than satisfactory specimen (e.g., insufficient amount, improper packaging, excessive delay in transport). Successful isolation of the pathogen requires that the best specimen be properly collected, promptly transported, and carefully processed. 21 1. Pulmonary Disease Although Mycobacterium tuberculosis is capable of causing dis- ease in almost any organ of the body, 85% of tuberculosis disease in the United States is still pulmonary (1, 21, 25,), as are the majority of diseases due to other mycobacteria (21, 27, 28). Thus, the majority of specimens submitted for diagnosis are lung secretions, obtained in several different ways. (1) (2) Sputum: To obtain a desirable sputum specimen, the patient should be instructed to: e Rinse the mouth with water before sputum is collected to minimize residual food particles, mouthwash, and oral drugs that might contaminate the specimen or inhibit growth of any acid-fast bacilli present e Remember that saliva and nasopharyngeal discharge are not sputum e Collect only the exudative material brought up from the lungs after a deep, productive cough Specimens should be collected in laboratory-approved containers, clearly labeled with patient name and/or identi- fication number. The number of specimens that should be submitted for culture is related to the results of early smear examination (1,16). When at least two of the first three sputum smears are positive, then three specimens usually are enough to confirm the diagnosis. If none, or only one of the first three sputum smears is positive, a larger number (usually 6) of specimens is needed for cultural confirma- tion of disease. A few patients shed mycobacteria in small numbers and irregularly; for these patients, the greater the number of specimens cultured, the greater the chance of obtaining a positive culture result (1). If a patient produces very little sputum, a 24- to 48-hour pooled specimen will often yield a positive culture (15). But if the specimen must be submitted by mail, the single, early morning collection is less frequently contaminated than the pooled specimen (13). If a patient finds it difficult to raise sputum, other methods may be used to obtain pulmonary secretions, as described below: Sputum Induction: The inhalation of warm, aerosolized hypertonic (5% to 10%) saline irritates the lungs enough to induce both coughing and the production of a thin, watery specimen (2, 5, 6, 11, 19). Because these specimens resem- ble saliva, itis very important that they be labeled “induced” specimens. 22 (3) (4) (5) Laryngeal Swabs (24, 26) may be useful in children and patients who raise no sputum or swallow it. Gastric Lavage should be considered when pulmonary secretions cannot be collected by the above methods. For optimum results, the patient should be hospitalized to col- lect the gastric lavage. The collection should be made early in the morning before the patient eats or gets out of bed. When sputum induction is followed in 30 minutes by gas- tric lavage, the combined results often yield more positive cultures than either method alone (5). Bronchial Washings, Biopsy, or Brushings and Transtracheal Aspiration not only produce a primary specimen, but the procedure causes the patient to produce sputum naturally for several days. 1. Extrapulmonary Disease Mycobacteria may not be suspected as the causative agent of an extrapulmonary disease because the chest roentgenogram is normal or the tuberculin skin test is negative, or both. Because mycobacteria may infect almost any organ in the body, the laboratory should expect to receive a variety of extrapulmonary specimens: aseptically col- lected body fluids, surgically excised tissues, aspirated or draining pus, and urine. These specimens may be divided into two groups (10): (1) Aseptically collected specimens, usually free of microorga- nisms (except the implicated pathogen) Specimens known to contain contaminating normal flora or specimens not collected aseptically Aspectically Collected Specimens — Fluid Body fluids (spinal, pleural, pericardial, synovial, ascitic, blood, pus, and bone marrow) are aseptically collected by the physician using aspiration techniques or surgical pro- cedure. Acid-fast bacilli may be difficult to isolate from some of these specimens because they often are diluted by the large fluid volume. Chances of isolating mycobacteria from these specimens can be improved if the laboratory person- nel and the clinician make previous arrangements to inocu- late aseptically collected fluid immediately into a liquid medium (Middlebrook 7H-9, Dubos Tween albumin broth, or Proskauer-Beck) at a ratio of 1 part fluid specimen to 5-10 parts liquid medium. Incubate cultures in the dark at 35 to 37°C in 10% CO, with daily shaking, by hand (do not use continuous agitation), to encourage the small numbers of mycobacteria to multiply. 23 (2) When weekly smear examinations of the inoculated broth reveal the presence of acid-fast bacilli, subculture the lig- uid medium to solid egg and/or agar base media for growth and identification of the organism. If the inoculated broth is still smear-negative after 4 to 6 weeks, centrifuge the entire volume (in aliquots, if necessary) at 3000 xg, or greater, and streak all the sediment to both egg and agar base media. When aseptically collected fluid that may clot cannot be inoculated immediately into a liquid medium, add sterile potassium oxalate (0.01 to 0.02 ml of 10% neutral oxalate solution per milliliter) or heparin (0.2 mg per milliliter) and transport the specimen to the laboratory as quickly as possible. Recovery of mycobacteria from blood, especially in cases of disseminated disease associated with acquired immuno- deficiency syndrome (AIDS), presents some unique prob- lems. Recently, the lysis-centifugation technique with the DuPont Isolator tube (13a) has successfully recovered M. chelonae (7a), M. avium complex, and M. tuberculosis bacilli from blood (Kiehn TE, personal communication). A 10-ml volume of blood is added to the Isolator tube; the tube is inverted several times to mix the contents, and then it is allowed to stand for 1 hour to lyse the blood cells. After lysis, the tube is centrifuged at 3000 x g or more to concen- trate the material into 1.5 to 2.0 ml of sediment, all of which is spread over the surface of several tubes or plates of isolation media (e.g., 7H-10, 7H-11, L-J). By using appropri- ate dilutions of sediment (or diluting lysed specimen prior to centrifugation), it is also possible to quantify the extent of the bacteremia. Aseptically Collected Specimens — Tissue Aseptically collected tissue specimens, suspected to con- tain mycobacteria, are placed in sterile containers without fixatives or preservatives. If the specimen is to be shipped by mail, protect the tissue from drying by adding sterile saline and pack the container in dry ice, or maintain a tem- perature of 5 to 10°C. If the specimen was indeed collected aseptically, homogenize it (only in a BSC) in a sterile tissue grinder (aerosol-free grinder preferred) with a small amount of sterile saline or sterile 0.2% bovine albumin, and inocu- late directly to liquid and solid media as with body fluids. When the tissue is not known to be sterile, homogenize it and inoculate half directly to liquid and solid media; the other half may be decontaminated as for sputum. Tissue material inoculated to liquid media should be examined 24 regularly by smear and inoculated onto solid media when smears become positive (as described above for sterile body fluids). (3) Specimens Expected to be Contaminated The majority of specimens received in the laboratory fall into this group, and most of them were discussed in the section on “Pulmonary Disease’; i.e., sputum, laryngeal swabs, bronchial washings, and gastric lavage. Among extrapulmonary specimens expected to be con- taminated are (a) those not handled aseptically after initial collection under sterile conditions and (b) clean-catch urine specimens from suspected genitourinary disease. In the former instance, handle specimens as you would sputum samples. Urine is the next most commonly encountered specimen that requires processing before culture. The genitourinary tract, one of the most common sites of extrapulmonary human tuberculosis (3, 22), accounts for nearly 20% of the extrapulmonary tuberculosis cases in the United States, and the incidence shows little evidence of decreasing (8). About 35% of the patients with genitourinary tubercu- losis also have tuberculosis elsewhere (3). Other mycobacteria act as noninvasive contaminants in the genitourinary tract and rarely as pathogens (27, 28). To minimize excessive contamination of urine specimens, the exter- nal genitalia should be washed before the specimens are collected, and the urine should be immediately processed or refrigerated. Either a single, early morning, voided midstream specimen (12) or the total first morning specimen (14) should be collected. Multiple, single speci- mens may be required to obtain positive results. From 50% to 70% of the cases are positive by smear examination with 25% to 95% posi- tive by culture (3, 18, 22). The AIDS patients with grossly disseminated mycobacterial dis- ease (e.g., M. avium complex and M. tuberculosis) occasionally pres- ent diagnostic problems that tax the ingenuity of laboratory personnel. Many specimens (e.g., lymph nodes, spinal fluids, tissue biopsies) are aseptically collected and may be handled as previously outlined. Some, such as blood, may be processed with technologies devel- oped in other areas of microbiology (see Isolator lysis-centrifugation method, page 24). Doubtless, new and innovative techniques will be spawned by the broadening demands of varied specimens from AIDS patients and other, as yet, unrecognized clinical conditions. B. Transportation The cetylpyridinium chloride (CPC) method (23) is used in some laboratories as a means of decontaminating sputum while in transit 25 -? to the laboratory. The use of this method not only decreases the number of cultures lost to contamination as a direct result of pro- longed in-transit time but also decreases significantly the laboratory time required for processing specimens (see “Isolation Procedures’’). The shipment of diagnostic specimens such as urine, sputum, and tissue, and of cultures of etiologic agents, which includes all Myco- bacterium spp., is subject to the packaging and labeling requirements of the Interstate Quarantine regulations (Federal Register, Title 42, Chapter 1, Part 72, revised July 30, 1972). Compliance with the require- ments of the regulation is the responsibility of the shipper. Failure to comply with the applicable requirements subjects violators to pen- alty provisions, which include fines and/or imprisonment. Diagnostic specimens must be packaged to withstand leakage of contents, shocks, pressure changes, and other conditions incident to ordinary handling in transportation. The “ETIOLOGIC AGENT/BIO- MEDICAL MATERIAL" label (described below) must not be affixed to shipments of diagnostic specimens. Etiologic agents are subject to additional specific containment pack- aging and labeling requirements. Cultures of Mycobacterium should be shipped to reference laboratories on solid medium in screwcap tubes. Petri dish cultures and cultures in liquid medium must not be shipped. Requirements for packaging etiologic agents are found in Section 72.25, paragraph (c) subparagraph (1) of the Interstate Quarantine regulations, as follows: Volume less than 50 ml. Material shall be placed in a securely closed, watertight container (primary container [test tube, vial, etc.]) which shall be enclosed in a second, durable watertight container (secondary container). Several primary contain- ers may be enclosed in a single secondary container, if the total volume of all the primary containers so enclosed does not exceed 50 ml. The space at the top, bottom, and sides between the primary and secondary containers shall contain sufficient nonparticulate material to absorb the entire contents of the primary container(s) in case of breakage or leakage. Each set of primary and secondary containers shall then be enclosed in an outer shipping container constructed of corrugated fiberboard, cardboard, wood, or other material of equiva- lent strength. See figure 9. Subparagraph (2) describes packaging of volumes of 50 ml or greater. This description is similar to subparagraph (1) except that the volume in a single primary container may not be over 500 ml and the total volume enclosed within the outer shipping container may not exceed 4000 ml. Subparagraph (3) states that if dry ice is used as a refrigerant, it must be placed between the secondary and outer container in such a way that, as the dry ice sublimates, the secondary container does not become loose inside the outer container. 26 LZ PRIMARY CONTAINER CULTURE ABSORBENT SPECIMEN RECORD (HSM 3.203) SHIPPING [ CONTAINER ADDRESS LABEL— WATER PROOF CROSS SECTION OF PROPER PACKING TAPE CULTURE ABSORBENT PACKING MATERIAL The Interstate Quarantine Regulations (Code of Federal Regula- tions, Title 42, Part 72.25, Etiologic Agents) was revised July 30, 1972, to provide for packaging and labeling requirements for etio- logic agents and certain other materials shipped in interstate traffic. The figure diagrams packaging and labeling of etiologic agents in volumes less than 50 ml in accordance with the provisions of sub- paragraph (C)(1) of the cited regulation. The Etiologic Agents-Biomedical Material label (see subparagraph (C)(4) of regulations) which must be placed on all shipments of etiologic agents is depicted below. The label must be 2 inches high by 4 inches long and printed in red ink on white stock. For further information on any provision of this regulation contact: Centers for Disease Control Attn: Office of Biosafety 1600 Clifton Road Atlanta, Georgia 30333 Telephone (404) 329-3883 ETIOLOGIC AGENTS oy BIOMEDICAL \ ) MATERIAL IN CASE OF DAMAGE OR LEAKAGE STANDARD FORM £20 JUNE 1973 | pRescrineo BY DEPT HEW (4 ? CFR) _420-101 NOTIFY DIRECTOR CDC ATLANTA, GEORGIA (404) 329-3311 FIGURE 9. Packaging and labeling of etiologic agents Department of Transportation regulations prohibit the shipment of more than 50 ml of an etiologic agent in a primary container on passenger aircraft. A volume greater than 50 ml may be carried by air freight provided it is properly packaged and the total volume within a single outer container does not exceed 4000 ml. An “ETIOLOGIC AGENT/BIOMEDICAL MATERIAL" label must be affixed to the outer container. These labels are available commer- cially (see figure 9); they should measure 2 inches high by 4 inches long, and be printed in red on a white background. Shippers of etio- logic agents must be familiar with the above regulations that govern packaging, labeling, and shipment of these products in interstate commerce. Questions regarding this regulation should be addressed to: Office of Biosafety Centers for Disease Control Atlanta, Georgia 30333 (Telephone 404/329-3883) 28 REFERENCES 7a. mn. 12. 13. 13a. 14. 15. 16. 17. 18. 19. 20. 21. . Bates JH. Diagnosis of tuberculosis. Chest 1979;76 (Supplement):757-63. . Beck GJ, Nanda K. Use of superheated aerosols as a diagnostic measure in tuberculosis. Dis Chest 1962;42:74-8. . Bentz RR, Dimcheff DG, Nemiroff MJ, Tsang A, Weg JG. The incidence of urine cultures positive for Mycobacterium tuberculosis in a general tuberculosis patient population. Am Rev Respir Dis 1975;111:647-50. . Blair EB, Brown GL, Tull AH. Computer files and analyses of laboratory data from tuberculosis patients. Il. Analyses of six years’ data on sputum specimens. Am Rev Respir Dis 1976;113:427-32. . Carr DT, Karlson AG, Stilwell GG. A comparison of cultures of induced sputum and gastric washings in the diagnosis of tuberculosis. Mayo Clin Proc 1967;42:23-5. . Elliott RC, Reichel J. The efficacy of sputum specimens obtained by nebulization versus gastric aspirates in the bacteriologic diagnosis of pulmonary tuberculosis. A comparative study. Am Rev Respir Dis 1963;88:223-7. . Engbaek HC, Weis-Bentzon M . Transport of sputum specimens to a central tuber- culosis laboratory. 1. Evaluation of routine specimens from Greenland. 2. Experi- mental work with sputum specimens from Denmark. Acta Tuberc Scand 1964; 45:89-104. Fajtasek MF, Kelly MT. Isolation of Mycobacterium chelonei with the lysis- centrifugation blood culture technique. J Clin Microbiol 1982;16:403-5. . Farer LS, Lowell AM, Meador MP. Extrapulmonary tuberculosis in the United States. Am J Epidemiol 1979;109:205-17. . Greenbaum M, Beyt BE Jr., Murray PR. The accuracy of diagnosing pulmonary tuberculosis at a teaching hospital. Am Rev Respir Dis 1980; 121:447-81. . Hawkins JE, Kubica GP, Wayne LG. Mycobacterium. In: Seligson D, ed.-in-chief. CRC handbook series in clinical laboratory science. Cleveland: CRC Press Inc., 1977;1:147-58. Jones FL Jr. The relative efficacy of spontaneous sputa, aerosol-induced sputa, and gastric aspirates in the bacteriologic diagnosis of pulmonary tuberculosis. Dis Chest 1966;50:403-8. Kenney M, Loechel AB, Lovelock FJ. Urine cultures in tuberculosis. Am Rev Respir Dis 1960;82:564-7. Kestle DG, Kubica GP. Sputum collection for cultivation of mycobacteria: an early morning specimen or the 24-72 hour pool? Am J Clin Pathol 1967;48:347-9. Kiehn TE, Wong B, Edwards FF, Armstrong D. Comparative recovery of bacteria and yeasts from lysis-centrifugation and conventional blood culture system. J Clin Microbiol 1983;19:300-4. Krasnow |. Primary isolation of mycobacteria. Lab Med 1978;9:26-31. Krasnow |, Wayne LG. Comparison of methods for tuberculosis bacteriology. Appl Microbiol 1969;18:915-7. Kubica GP, David HL. The mycobacteria. In: Sonnenwirth AC, Jarett L, eds. Gradwohl’s clinical laboratory methods and diagnosis. 8th ed. St. Louis: C.V. Mosby Co., 1980:1693-730. Kubica GP, Gross WM, Hawkins JE, Sommers HM, Vestal AL, Wayne LG. Labora- tory services for mycobacterial diseases. Am Rev Respir Dis 1975;112:773-87. Lattimer JK. Renal tuberculosis. N Engl J Med 1965;273:208-11. Lillehei JP. Sputum induction with heated aerosol inhalations for the diagnosis of tuberculosis. Am Rev Respir Dis 1961;84:276-8. MacGregor RR. A year’s experience with tuberculosis in a private urban teaching hospital in the postsanatorium era. Am J Med 1975; 58:221-8. Ortbals DW, Marr JJ. A comparative study of tuberculosis and other mycobacterial infections and their association with malignancy. Am Rev Respir Dis 1978;117:39-45. 29 22. 23. 24. 25. 26. 27. 28. Simon HB, Weinstein AJ, Pasternak MS, Swartz MN, Kunz LJ. Genitourinary tuberculosis. Clinical features in a general hospital. Am J Med 1977;63:410-20. Smithwick RW, Stratigos CB, David HL. Use of cetylpyridinium chloride and sodium chloride for the decontamination of sputum specimens that are transported to the laboratory for the isolation of Mycobacterium tuberculosis. J Clin Microbiol 1975;1:411-3. Tonge JI, Hughes PG. A comparative study of laryngeal swabs and gastric lavage in the detection of tubercle bacilli. Am Rev Tuberc 1956;73:930-9. Tuberculosis in the United States, 1978. Atlanta: Centers for Disease Control, PHS, (HHS, 1980 HHS Publications No. (CDC) 80-8322). Velu S, Narayana SSL, Subbaiah TV. A comparison of the results of bacteriologic examination of a sputum collection and a pair of laryngeal swab specimens in patients receiving chemotherapy for pulmonary tuberculosis. Tubercle 1962;43:1-10. Wolinsky, E. State of the art: nontuberculous mycobacteria and associated diseases. Am Rev Respir Dis 1979;119:107-59. Wolinsky E. The impact; clinical and epidemiologic observations: human diseases. In: Kubica GP, Wayne LG, Good LS, eds. 1954-1979: Twenty-five years of mycobacte- rial taxonomy. Atlanta: Centers for Disease Control, PHS, HHS, 1980:13-8. 30 Isolation Procedures Isolation Procedures Specimens collected aseptically (surgically excised tissue, aspira- tions from closed lesions, and sterile body fluids) from patients sus- pected of having a mycobacterial disease may be inoculated directly onto primary isolation media without prior decontamination, pro- vided they are properly handled and homogenized before inoculation. Unfortunately, the majority of clinical specimens submitted to the mycobacteriology laboratory for cultural confirmation of the suspected etiologic agent are contaminated to varying degrees by more rapidly growing normal flora organisms. Most of these specimens must be subjected to a harsh digestion-decontamination procedure that lique- fies the organic debris and eliminates the unwanted normal flora organisms that would otherwise overgrow the more slowly growing mycobacteria on the culture media. All currently available digesting/ decontaminating agents are toxic (at least to some extent) for both the mycobacteria and the more rapidly growing contaminants; therefore, to ensure the survival of maximal numbers of mycobacteria in the submitted specimen, the digestion procedure must be pre- cisely followed. The success of any of these procedures also depends on: the relatively greater resistance of the mycobacteria to strongly alkaline or acidic digesting solutions; the length of time the mycobac- teria are in contact with the digesting-decontaminating agent; the temperature buildup in the specimen during centrifugation; and the efficiency of the centrifuge used to sediment the mycobacteria. The digestants most commonly used in the United States for processing of clinical specimens are: N-acetyl-L-cysteine-(NALC) sodium hydroxide (5) Zephiran-trisodium phosphate (Z-TSP) (12) Sodium hydroxide (NaOH) (8) A. Centrifugal Efficiency and Digestant Toxicity Most early reports (and many recent ones) describing sputum pro- cessing in the tuberculosis (TB) laboratory recorded centrifuge speeds in revolutions per minute (rpm). However, revolutions-per-minute is a measure of speed for a particular centrifuge head and not a mea- sure of sedimenting efficiency or relative centrifugal force (RCF). The latter may be calculated from the formula, RCF = 1.12 R max (rpm/ 1000)%, where R max = radius (mm) from the center of the rotating head to the bottom of the spinning centrifuge tube. If appropriate 31 equipment is available, RCF may be reproduced in different labora- tories. For example, the required rpm to generate a desired RCF may be calculated from the following formula: RCF rom = 1000 erreeme 1.12 R max Recent studies on the sputum digestion process have addressed the toxicity of the digestant-decontaminant (4,7), centrifugal efficiency (7,9), heat buildup during centrifugation and its effect on the contin- ued killing action of the digestant (7), and contribution of all the foregoing to the overall “toxicity” of the digestion-decontamination process. Several investigations (4,7) have provided confirmatory data that the NALC-NaOH and Z-TSP digestion-decontamination procedures result in the death of 28% to 33% of the mycobacteria in a clinical specimen, whereas the 4% NaOH method of Petroff kills 60% to 70% of the mycobacteria. This initial kill by the digestant is independent of the additional contributory factors of heat buildup in the centrifuge and cenrifugal efficiency. Recent studies of Rickman and Moyer (9) alerted us to the sedi- menting efficiency of higher RCF and demonstrated the painful fact that most mycobacteriology laboratories have done a very poor job of sedimenting mycobacteria from digested sputum specimens. Our own studies (7) confirmed that centrifuges used in the “TB Labora- tory” were very inefficient in terms of sedimenting the acid-fast bacilli. If RCF is not high enough, many mycobacteria remain in suspension following centrifugation and are poured off with the discarded super- natant fluid. Figure 10 shows the log of the percentage of cells that still remain in the supernatent fluid following centrifugation at selected RCF or gravities (g-forces) used in many clinical laboratories. On the basis of the observed percentages of mycobacteria that were left in suspension following centrifugation at a number of RCF-time com- binations, it was possible to construct a formula that provides a fairly reliable measure of centrifugal efficiency: Log10% AFB in suspension = 2.6371-0.0426 (time)-0.0004 (RCF). Subtraction from 100 of the calculated percentage still in suspen- sion gives the sedimenting efficiency (or percentage of cells sedi- mented) at any RCF-time combination from 1000 to 4000 x g. Studies in this laboratory (7) suggest that a 95% sedimenting efficiency should be sought for all specimens processed in the mycobacteriology laboratory. Many of the old centrifuges still used in mycobacteriology labora- tories (8-, 12-, and 16-place horizontal heads) commonly spin at 2300 to 3000 rpm (only 1500 to 2000 RCF); most users of such equipment spin their digested specimens for only 15 minutes and thus attain 32 theoretical sedimenting efficiencies ranging from 75%-84% (in actual practice, heat buildup during centrifugation reduces efficiency still more; see below). The sedimenting efficiency of lower g-forces (RCF) may be made to equal that of higher g-forces by extending the time of centrifugation to satisfy the formula t;RCF,; = t,RCF,, where RCF’'s 1 and 2 are the g-forces, and t's 1 and 2 are the times required to make the two forces yield comparable sedimenting efficiency. Because lower g-forces will Log, o Bacilli Still in Suspension Time (in minutes) FIGURE 10. Efficiency of centrifugation Log; of percentage of acid-fast bacilli in suspension vs. centrifugation time and g-forces. Broken line indicates 95% sedimentation of bacilli (or 5% still in suspension, i.e., Log 0.7). 33 require longer times to effect sedimentation comparable to higher g-forces, it is important to know (a) the lethal effect on mycobacteria of longer contact with the digestant and (b) the possible increase in this lethal effect due to frictional temperature elevation in the rapidly spinning centrifuge tubes (7,14). Depending on the number of replicate runs and the type of rotor used in a nonrefrigerated centrifuge, the temperature increase within the specimen tube may vary from 4 to 18°C above ambient (7). Tubes spun in a streamlined angle head are least affected by temperature rise even after several runs, but the contents of tubes spun in unpro- tected horizontal rotors may exceed 40°C if the centrifuge has been used for 5 or 6 successive runs. Figure 11 indicates the lethal effect on mycobacteria of the diluted NALC-NaOH digestant (just before centrifugation) that results from the combined influence of tempera- ture elevation and centrifuge spinning time. Note that even when centrifuge time remains constant at 15 minutes, the percentage of cells killed increases from 13% to 22% to 30% as the temperature rises from 20 to 30 to 40°C. It seems important, therefore, to keep the spinning time low (15 minutes) and the RCF high (3000x g) to effect 95% sedimentation. Too, the use of angle head rotors minimizes heat buildup due to air friction. Our studies (7) suggest that refrigeration of the centrifuged specimen at 8 to 10°C might eliminate the lethal effect of heat on mycobacteria, but streamlined rotors and short time, high- speed runs in unrefrigerated centrifuges would work almost as well. 13) | 15 min o 20 C 17) | 20 min 30 min 15 min o 30 C 20 min o 40 C 30 min FIGURE 11. Combined lethal effect on mycobacteria of temperature and time of exposure to a potentially lethal digestant 34 Doubtless, most other sputum digestants exhibit a time-temperature related toxicity for mycobacteria. Figure 12 illustrates the overall kill of mycobacteria that resulted from the use of three different methods for digestion and decontami- nation of sputum specimens that have been studied or used at CDC over the years. Results of recent studies emphasize that for optimum recovery of mycobacteria from clinical specimens: 1. The digestion-decontamination procedure should be as gen- tle as possible (compatible with an overall contamination rate not in excess of 5%) 2. The centrifuge rotor should be one that minimizes frictional air resistance (i.e., angle head, or, possibly, a windscreen bowl) 3. The RCF should be as high as compatible with centrifuge tubes in use* 4. The spin time should be of duration to provide 95% sedimen- tation with minimal loss of cells due to heat build up.* + NALC- NaOH 42% (3000g/15 min) 3 J 40% (1 wash, 1 spin at 2000g/20 min) Z-TSP 46% (2 wash, 2 spin at 2000g/20 min) 4% NaOH 1)+(2 73% (14009/15 min) Cumulative loss indicated by numbered circles: 1 = initial kill by digestant-decontaminant. 2 = continued killing during centrifugation. 3 = efficiency of centrifugation, i.e., cells not sedimented. FIGURE 12. Hypothetical examples of overall kill with different di- gestants * The tolerance of 50-ml conical bottom centrifuge tubes for g-forces in excess of 3000 RCF varies with the manufacturer (check supplier for this information). The 50-ml conical bottom centrifuge tubes that were first made for use in the tuberculosis laboratory could tolerate only about 3200 g; more recently, tubes that can withstand 5000 g have been manufactured. * + Our recent studies (7) showed that 3000 g for 15 minutes would sediment 95% of the mycobacteria in a digested sputum specimen. From the for- mula on page 33, it is interesting to speculate that only 5 minutes of centrifugation in new tubes that can tolerate 5000 g would theoretically sediment 95% to 97% of suspended bacilli. Studies to confirm these theo- retical figures have not yet been performed. 35 B. Digestion-Decontamination Procedures 1. N-Acetyl-L-Cysteine-Sodium Hydroxide (NALC-NaOH) Method (5,6,10) The mucolytic agent, N-acetyl-L-cysteine, used for rapid digestion of sputum, enables the decontaminating agent, sodium hydroxide, to be used at a lower final concentration (in sputum) of 1%. Acetylcysteine loses activity rapidly in solution, so the digestant should be made fresh daily. Sodium citrate is included in the digestant mixture to bind the heavy metal ions that might be present in the specimen and could inactivate the acetylcysteine (6). eo Materials N-acetyl-L-cysteine-NaOH (NALC-NaOH) solution Centrifuge tubes, 50-ml Media: Agar base such as Middlebrook 7H-10 or 7H-11 Egg base such as Lowenstein-Jensen (LJ) or American Trudeau Society (ATS) Microscope slides 0.067M Phosphate buffer, pH 6.8 OR Water, sterile distilled, 30 to 40 ml per specimen Water, sterile distilled, 4.5 ml for dilution blanks Bovine albumin, 0.2% in saline. e Preparations (1) The NaOH and sodium citrate may be mixed (table 1), sterilized, and stored in sterile screwcap flasks for later use. After NALC has been added, the prepared volume of digestant must be used within 24 hours, because NALC loses mucolytic activity on standing. TABLE 1. Preparation of NALC-NaOH digestant-decontaminant solution. Volume of Mix indicated amounts (ml) of Add NALC Digestant (grams) Needed (ml) 4% NaOH(*) 2.9% Na citrate 2H,0( +) 50 25 25 0.25 100 50 50 0.50 200 100 100 1.00 500 250 250 2.50 1000 500 500 5.00 (*) Add 4.0g NaOH to 100 ml distilled water. (+) Add 2.9g sodium citrate dihydrate (or 2.6 g anhydrous sodium citrate) to 100 ml distilled water. 36 (2) (3) (4) 0.067M Phosphate buffer, pH 6.8 Stock solutions (a) Disodium phosphate - dissolve 9.47 g of anhydrous Na,HPO, in 1 liter of dis- tilled water. (b) Monopotassium phosphate - dissolve 9.07 g of KH,PO, in 1 liter of distilled water. To prepare pH 6.8 buffer solution, mix 50 ml (a) with 50 ml (b) and check pH on meter. If final buffer requires pH adjustment, add solution (a) to raise the pH or solution (b) to lower it. Bovine albumin, 0.2% 2.0% Stock solution: Dissolve 0.85 g NaCl in 100 ml of dis- tilled water, add and dissolve 2 g of bovine albumin fraction V by swirling or stirring gently on a magnetic stir- rer. Adjust pH to 6.8 with 4% NaOH and sterilize the solution by Seitz or membrane filtration. 0.2% Use solution: Dilute 2% stock solution 1:10 with sterile 0.85% saline. Label all media, slides, and tubes for proper patient identifi- cation and date of inoculation or preparation. Procedure for Sputum (1) (2) (3) (4) Work in sets equivalent to one centrifuge load (e.g., 8 speci- mens at at time). Transfer 10 ml, or less, of sputum to 50-ml plastic centrifuge tube and add an equal volume of NALC-NaOH solution. After digestant is added to one set of tubes, tighten caps of tubes and mix on test tube mixer until liquefied (about 5 to 20 seconds per tube) and invert each tube to insure that NALC-NaOH solution contacts all inside surfaces of the tube and caps. Caution: Avoid extreme agitation to minimize oxidation and inactivation of the acetylcysteine. Centrifugal mixing of most test tube mixers provides the desired kind of agitation. Let tubes stand 15 minutes at room temperature for decon- tamination. If more active decontamination is needed, increase concentration of NaOH in table 1 to 5% or 6% rather than increasing the time the specimen is exposed to digestant. Dilute the digested-decontaminated specimen to the 50 ml mark with sterile distilled water or sterile phosphate buffer, pH 6.8, to minimize the continuing action of NaOH and to 37 (5) (6) (8) (9) lower the specific gravity of the specimen before centrifug- ing. Tighten tube caps and mix by swirling or inversion. Centrifuge at 3000 x g for 15 minutes (or the appropriate RCF-time combination to give 95% sedimentation) using aerosol-free sealed centrifuge cups. If aerosol-free cups are not available, the centrifuge may be modified to control aerosols, as described by Hall (2). After centrifugation, pour off supernatant fluid into a splash- proof discard can containing disinfectant, swab lip of tube with disinfectant soaked gauze (or flame carefully), and recap. Resuspend sediment in 1 to 2 ml of sterile 0.2% bovine albumin solution. If media will be inoculated immediately, the sediment may be resuspended in sterile saline or sterile water. Mix sediment with 1-ml pipette and make a 1:10 dilution of the resuspended sediment (see figure 13) to decrease the concentration of any toxic substances that may inhibit growth of mycobacteria. 10m 170ml 1.0m! 1.0ml 1.0ml CNC NTN ON broth culture 10-3 10-4 10~° 1:1000 1:10,000 1:100,000 10-2 1:100 10-1 1:10 (10) (11) FIGURE 13. Diagram for tenfold dilutions Inoculate 0.1 ml of both the undiluted and 10" dilution of the digest onto each of two tubes of an egg base (L-J) and one half of a biplate of an agar base (7H-10 plate) medium. Make a smear of the undiluted sediment by spreading a drop over an area 1 x 2 cm on the microscope slide. Allow to air dry; heat-fix at 65 to 75°C for 2 hours, or overnight (or pass slide 3 to 4 times through the blue cone of a burner flame); stain and examine (see ‘‘Acid-Fast Microscopy’ for details). 38 (12) (13) (14) Place 7H-10 plates in individual CO,-permeable polyethyl- ene bags and incubate at 35 to 37°C under 10% CO in air. Incubate L-J tubes at 35 to 37°C in a horizontal position, making certain caps are loose. An atmosphere of 10% CO, is not required but will encourage earlier, more luxuriant growth. Refrigerate the remaining undiluted sediment for later use if direct drug tests may be requested. Procedure for Gastric Lavage These specimens should be processed within 4 hours of collec- tion. (1) (2) (3) (4) (5) If specimen is quite watery, proceed to step 3. If specimen is mucoid, (a) add 50 to 100 mg of NALC pow- der to 50 ml of gastric lavage, replace and tighten the cap; (b) mix by swirling on a test tube mixer. Centrifuge at 3000 x g for 15 minutes in aerosol-free safety carriers. After centrifugation, pour off supernatant fluid and resus- pend sediment in 2 to 5 ml of sterile distilled water. Add an equal volume of NALC-NaOH and proceed as for sputum. Procedure for Laryngeal Swab (1) (2) (3) (4) (5) (6) (7) Use sterile forceps to transfer the swab to a sterile centri- fuge tube. If necessary, break off the swab so the tube cap can be replaced. Add 2 ml of sterile distilled water or saline. Add 2 ml of NALC-NaOH. Swirl on test tube mixer. Let stand 15 minutes to decontaminate. Remove swab from centrifuge tube using sterile forceps. Fill the tube with sterile buffer or water and proceed as for sputum. Procedure for Tissue (1) (2) Use a sterile tissue grinder (aerosol-free preferred) and ster- ile 0.85% saline or 0.2% bovine albumin to grind tissues. If tissue (lung) contains mucus, add a pinch of NALC. If tissue has been collected and processed aseptically, inocu- late the homogenate directly to both liquid and solid media. Using aseptic technique, make weekly smears of broth culture, stain, and examine for acid-fast bacilli. (a) When smear is positive for AFB, inoculate the broth onto solid media (egg and agar). (b) If, after 4 weeks, smears remain negative, centrifuge total volume of liquid medium at 3000x g (or more) for 15 minutes. Place the tip of a 1-ml pipette at the 39 (3) very bottom of the tube and remove 0.5 to 1.0 ml of fluid; inoculate to multiple egg slants and agar plates. Incubate for 4 to 8 weeks at 35 to 37°C (also at 30 to 32°C, if lesion was superficial). If tissue has not been aseptically collected and handled (or, if there is doubt), a portion of the homogenate may be inoculated directly to media while the remainder is treated as for sputum. Procedure for Blood (1 (2) (3) (4) (5) Add 10 ml of aseptically collected blood to 50 ml of a Tween- containing liquid medium such as Dubos Tween albumin broth or Middlebrook 7H-9. Several tubes or plates of solid media (e.g., LJ or 7H-11) may also be inoculated with the specimen. Incubate in the dark at 35 to 37°C. Shake liquid cultures once daily by hand. Prepare smears of liquid cultures once a week. Stain and examine smears under the microscope (fluorescence mi- croscopy, is easier to read) for presence of acid-fast bacilli. Inoculate solid media with any smear-positive liquid cul- tures. Incubate solid media for 4 to 8 weeks before discarding as negative. More recently, the Dupont Isolator lysis-centrifu- gation tube has been used with great success to recover several different species of Mycobacterium from blood (see details, p. 24). Procedure for Other Body Fluids MUCOPURULENT MATERIALS: (1) (2) Handle as for sputum when volume is 10 ml or less. Handle as for mucoid gastric lavage when volume is more than 10 ml. FLUID MATERIALS: (1) (2) If collected aseptically, centrifuge and inoculate sediment directly onto culture media. If not aseptically collected: a. Handle as for sputum when volume is 10 ml or less. b. Handle as for fluid gastric lavage when volume is more than 10 ml. 2. Zephiran - Trisodium Phosphate (Z-TSP) Method (10, 12) The use of trisodium phosphate and Zephiran (benzalkonium chloride) to homogenize and decontaminate specimens is another of the more gentle digestion procedures. Since Zephiran is bacterio- static to mycobacteria, the digested, centrifuged sediment must be 40 neutralized by washing with buffer before inoculating onto agar medium (7H-10 or 7H-11). The phospholipids of egg medium provide a “built-in”’ neutralizer for this quaternary ammonium compound, making neutralization unnecessary (3). Materials Zephiran-trisodium phosphate (Z-TSP) solution Neutralizing buffer, pH 6.6 Centrifuge tubes, 50-ml Media: Agar base - 7H-10 or 7H-11 Egg base - L-J, ATS Microscope slides Preparations (1 (2) Z-TSP Solution Dissolve 1 kg of trisodium phosphate (Na3PO4:12 H;0) in 4 liters of hot distilled water. Add 7.5 ml of Zephiran (17% benzalkonium chloride, Win- trop Laboratories). Mix well and store at room temperature. Neutralizing buffer, pH 6.6 Mix 37.5 ml of disodium phosphate buffer (a) and 62.5 ml monopotassium phosphate buffer (b) (for preparation of buffer solutions, see page 37). Check pH on meter. Bottle in 20-ml| amounts, autoclave, and store at room temperature. Procedure (1) (2) (3) (4) (5) (6) (7) (8) (9) Transfer 10 ml, or less, of sputum to 50-ml plastic centri- fuge tube and add an equal volume of Z-TSP solution. Tighten cap of tube and agitate vigorously on a mechanical shaker for 30 minutes. Let stand 20-30 minutes without additional shaking. Centrifuge at 3000 x g for 15 minutes. After centrifugation, decant the supernatant fluid and resus- pend the sediment in 20 ml of neutralizing buffer, pH 6.6, and mix well. Centrifuge as before for 15 minutes. Decant and discard the supernatant fluid, retaining some fluid to resuspend the sediment. Mix the sediment with a pipette and inoculate three drops to each tube (L-J) and plate (7H-10) of medium. Incubate egg medium horizontally at 35 to 37°C with caps loosened. Place agar plates in plastic bags and incubate in 10% CO, in air at 35 to 37°C. a1 (10) Make a smear of the sediment. Allow to air dry, heat fix, stain, and examine. 3. Petroff’s Sodium Hydroxide (NaOH) Method (8) NaOH is toxic, not only for the non-acid-fast contaminants but also for mycobacteria; therefore, during this digestion procedure adhere strictly to the indicated timing. The digestant, 2% to 4% NaOH, should be used at the lowest concentration that will effectively digest and decontaminate the specimen. eo Materials NaOH Solution (2% to 4%) 2N HCI solution or HCI-phenol red indicator e Preparations (1) NaOH Dissolve NaOH in distilled water to give desired concentra- tion (2 g/100 ml for 2% or 4 g/100 ml for 4%). Sterilize by autoclaving. (2) 2N HCI Solution Dilute 33 ml of concentrated HCI to 200 ml with water (always add acid to water, never the reverse). Sterilize by autoclaving. (3) Phenol red indicator Combine 20 ml of phenol red solution (0.4% in 4% NaOH) and 85 ml of concentrated HCI with distilled water to make 1000 ml. Note, the phenol red solution may also be used separately. e Procedure (1) To 10 ml of specimen in a 50-m| screwcapped tube, add equal volume of appropriate NaOH solution. (2) Shake vigorously on mechanical mixer (e.g., paint condi- tioning machine) for 15 minutes or, preferably, mix on test tube mixer to digest and let stand for 15 minutes. (3) Centrifuge at 3000 x g for 15 minutes. (4) After sedimentation, decant supernatant fluid and neutral- ize sediment with HCI. Either add one drop indicator solu- tion followed by adding HCI dropwise, or use HCl-phenol red indicator. Add HCI until indicator changes from red to persistent yellow. (6) Mix sediment to resuspend and inoculate desired media. (6) Make a smear, allow to air dry, heat fix, stain and examine. With any digestion-decontamination procedure, a contamination rate of 5% is acceptable. If the rate of contamination varies markedly 42 from this, the digestion procedure should be examined. Over-digestion of the specimens would cause the percentage of contamination to be consistently lower than the “acceptable” 5%, whereas under-digestion would yield a higher percentage of contaminated cultures. When either situation occurs with alkaline digestion-decontamination solutions, it is better to alter the concentration of the alkali in the initial digestant solution rather than to increase or decrease the digestion time for the specimen. If a particular patient's specimens are consistently contami- nated (indicating that the routine digestion-decontamination proce- dure is unsatisfactory), a special procedure might be necessary. A few such procedures are listed below. 4. Oxalic Acid Method (1) This method is often helpful with those specimens consistently contaminated with Pseudomonas species. eo Materials 5% Oxalic acid 4% Sodium hydroxide Physiological saline (0.85%) Phenol red indicator e Preparations (1) 5% Oxalic acid Dissolve 5 g oxalic acid in 100 ml of distilled water and autoclave to sterilize. (2) 4% Sodium hydroxide Dissolve 4 g NaOH in 100 ml of distilled water and auto- clave to sterilize. (3) Physiological saline Dissolve 0.85 g NaCl in 100 ml of distilled water and auto- clave to sterilize. (4) Phenol red indicator Dissolve 8 mg of phenol red powder in 20 ml of 4% NaOH and add sufficient distilled water to make 1000 ml. e Procedure (1) Add an equal volume of 5% oxalic acid to 10 ml, or less, of specimen in a 50-ml centrifuge tube. (2) Vortex and let stand 30 minutes with occasional shaking. (3) Add sterile saline to the 50 ml mark on the centrifuge tube. (4) Centrifuge for 15 minutes at 3000 x g. (5) Decant the supernatant fluid and add a few drops of phenol red indicator to the sediment. (6) Neutralize with 4% NaOH until a persistent pale pink color forms. 43 (7) Resuspend the sediment, inoculate onto media, and incu- bate at 35 to 37°C. Make smears, air dry, heat fix, stain, and examine. 5. Sulfuric Acid Method (13) This method is sometimes helpful for urine and other thin watery body fluids that consistently yield contaminated cultures when pro- cessed with one of the alkaline digestants. e Materials 4% Sulfuric acid 4% Sodium hydroxide Sterile distilled water Phenol red indicator eo Preparations (1) 4% Sulfuric acid Slowly add 40 ml of concentrated sulfuric acid to 960 ml of distilled water. Store at 4°C. (2) 4% Sodium hydroxide Dissolve 4 g NaOH in 100 ml of distilled water and auto- clave to sterilize. (3) Phenol red indicator Dissolve 80 mg of phenol red powder in 20 ml of 4% NaOH and add sufficient distilled water to make 1000 ml. e Procedure (1) Centrifuge entire specimen for 30 minutes at 3000 x g. This may require several tubes. (2) Decant supernatant fluid and pool sediments if several centrifuge tubes were used for a single specimen. (3) Add an equal volume of 4% sulfuric acid to the sediment. (4) Mix on test tube mixer and let stand for 15 minutes at room temperature. (5) Fill the tube to the 50-mI| mark with sterile water. (6) Centrifuge at 3000 x g for 15 minutes. (7) Decant supernatant fluid. (8) Add one drop phenol red indicator and neutralize with 4% NaOH until a persistent pale pink color forms. (9) Inoculate desired media and incubate at 35 to 37°C. (10) Make smears of sediment and stain for AFB. 6. Cetylpyridinium Chloride-Sodium Chloride (CPC) Method (11) The CPC method was proposed as a means of digesting and decon- taminating sputa in transit. After it arrives in the laboratory, the 44 digested-decontaminated specimen need only be concentrated by centrifugation and the sediment inoculated directly onto media. This method, of course, provides a significant decrease in time required to process sputum specimens. This procedure provided an increase in the number of cultures positive for mycobacteria other than M. tuber- culosis and resulted in less contamination than the NALC-NaOH method. Cetylpyridinium chloride, a quaternary ammonium compound, is used to decontaminate the specimen while sodium chloride effects liquefaction. Since CPC is bacteriostatic for mycobacteria inoculated onto agar base medium and this effect is not neutralized in the diges- tion procedure, sediments from specimens treated with CPC should be inoculated only onto egg base medium. e Materials 1% Cetylpyridinium chloride 2% Sodium chloride Sterile water, 20 to 40 ml per specimen Sterile saline, or 0.2% bovine albumin fraction V. e Preparation (1) CPC Digestant-Decontaminant Dissolve 10 g of cetylpyridinium chloride and 20 g of NaCl in 1000 ml of distilled water. The solution is self- sterilizing and remains stable for an extended period if pro- tected from light, extreme heat, and evaporation. Dissolve with gentle heat any crystals that might form in the work- ing solution. (2) 0.2% Bovine albumin fraction V 2% Stock solution—Dissolve 0.85 g of NaCl in 100 ml of distilled water; add 2 g of bovine albu- min fraction V and solubilize by pro- longed stirring. Adjust pH to 6.8 with 4% NaOH. Sterilize by Seitz or mem- brane filtration. 0.2% Use solution—Dilute 2% stock solution 1:10 sterile saline. e Procedure (1) Collect 10 ml, or less, of sputum in 50-m| screwcap centri- fuge tube. (2) Add an equal volume of CPC-NaCl, cap securely, and shake by hand (inside biological safety cabinet) until specimen liquefies. (3) Package specimen in double mailing container and send to processing laboratory. The specimen is digested and decon- taminated in transit. 45 REFERENCES 10. 11. 12. 13. 14. (4) Upon receipt in the laboratory, dilute the digested-decon- taminated specimen to the 50-m| mark with sterile distilled water, replace, and tighten cap securely. (6) Centrifuge at 3000 x g for 15 minutes. (6) Decant supernatant fluid and suspend sediment in 1 to 2 ml of sterile water, saline, or 0.2% bovine albumin fraction V. (7) Inoculate resuspended sediment onto egg medium and incubate at 35 to 37°C. . Corper HJ, Uyei N. Oxalic acid as a reagent for isolating tubercle bacilli and a study of the growth of acid fast nonpathogens on different mediums with their reactions to chemical reagents. J Lab Clin Med 1930; 15:348-69. . Hall CV. A biological safety centrifuge. Health Lab Sci 1975;12:104-6. . Krasnow |, Kidd GC. The effect of a buffer wash on sputum sediments digested with Zephiran trisodium phosphate on the recovery of acid-fast bacilli. Am J Clin Pathol 1965;44:238-40. . Krasnow |, Wayne LG. Sputum digestion. |. The mortality rate of tubercle bacilli in various digestion systems. Am J Clin Pathol 1966;45:352-5. . Kubica GP, Dye WE, Cohn ML, Middlebrook G . Sputum digestion and decontami- nation with N-acetyl-L-cysteine-sodium hydroxide for culture of mycobacteria. Am Rev Respir Dis 1963;87:775-9. . Kubica GP, Kaufman AJ, Dye WE . Comments on the use of the new mucolytic agent, N-acetyl-L-cysteine, as a sputum digestant for the isolation of mycobacteria. Am Rev Respir Dis 1964;89:284-6. . Kubica GP, Kent PT. The sputum digestion process in the mycobacteriology laboratory: contributions of centrifugal-efficiency and digestant toxicity. (In preparation, 1985.) . Petroff SA. A new and rapid method for the isolation and cultivation of tubercle bacilli directly from the sputum and feces. J Exp Med 1915;21:38-42. . Rickman TW, Moyer NP. Increased sensitivity of acid-fast smears. J Clin Microbiol 1980;11:618-20. Runyon EH, Karlson AG, Kubica GP, Wayne LG. Mycobacterium. In: Lennette EH, Spaulding EH, Truant JP, (eds.) Manual of clinical microbology. 2nd ed. Washington: American Society for Microbiology, 1974;150. Smithwick RW, Stratigos CB, David HL. Use of cetylpyridinium chloride and sodium chloride for the decontamination of sputum specimens that are transported to the laboratory for the isolation of Mycobacterium tuberculosis. J Clin Microbiol 1975;1:411-3. Wayne LG, Krasnow |, Kidd GC. Finding the “hidden positive” in tuberculosis eradication programs. The role of sensitive trisodium phosphate-benzalkonium (Zephiran) culture technique. Am Rev Respir Dis 1962;86:537-41. Willis HS, Cummings MM. Diagnostic and experimental methods in tuberculosis. 2nd ed. Springfield, Ill: Charles C Thomas, 1952. Working Party, Public Health Laboratory Service on Laboratory Diagnosis of Tuberculosis (Phease, RN, Chairman). a. Cultivaton of Mycobacterium tuberculo- sis. Il. Temperature changes during centrifugation. Month Bull Minist Health. 1953; 12:232-7. b. lll. The influence of time of centrifugation on cultural results. Month Bull Minist Health 1953;12:238-42. 46 C. Culture Media The definitive diagnosis of mycobacterial disease demands that the causative agent be recovered on culture medium and identified by using a number of differential in vitro tests. Many different media are available for cultivating mycobacteria; most are variations of either egg-potato base or serum (albumin) -agar base media. Because there is no general agreement on which medium is best for the routine isolation of acid-fast organisms, culture medium usually is selected on the basis of personal preference and/or laboratory tradition. The ideal medium for isolation of mycobacteria should (a) support rapid and luxuriant growth of small numbers of mycobacteria, (b) permit preliminary differentiation of isolates on the basis of pigment produc- tion and colony morphology, (c) inhibit the growth of contaminants, (d) be economical and simple to prepare from readily available ingredients, and (e) enable the performance of drug susceptibility tests. Since no single medium meets all these requirements and because our own experience has shown some isolates to grow on one medium and not another, we recommend that both an agar-base (e.g., 7H-10 or 7H-11) and an egg-base (e.g., Lowenstein-Jensen or American Trudeau Society) medium be used for primary isolation of mycobacteria. Using media of two different basal compositions enables the isolation of most strains of mycobacteria that cause human disease (14). 1. Advantages and Disadvantages of Egg-Base vs. Agar-Base Media e The advantages of an egg-base medium are: (1) It may be stored in the refrigerator for several months pro- vided it was made from fresh eggs and if the caps are tightly closed to minimize drying by evaporation. (2) Itis less likely to become contaminated during preparation because it is inspissated after being placed in tubes. (3) It supports good growth of most mycobacteria. e The disadvantages of an egg-base medium are: (1) When contamination does occur, it usually involves the total surface of the medium. (2) Drug susceptibility tests are more difficult to perform on egg media because the concentrations of certain drugs must be adjusted to account for their loss by heating or by inter- action with certain components of the egg (e.g., phos- pholipids). e The advantages of an agar-base medium are: (1) Itis less likely to become contaminated because its simpler chemical formulation is less likely to support contaminants. 47 (2) Itis usually a clear medium that enables the earlier observa- tion of colony morphology because, in contrast to opaque egg medium, the light-transparent agar medium may be examined with the aid of a dissecting microscope (30-60X). (3) Drug susceptibility tests may be performed with more pre- cise concentrations of desired drugs, because the medium has a simpler formulation than egg media and it is solidi- fied by agar rather than by heat. (4) If incubated in an atmosphere of 10% C0,-90% air, 99% of the cultures inoculated onto agar media (7H-10 or 7H-11) are positive within 3 to 4 weeks. e The disadvantages of the agar-base medium are: (1) Medium contained in plates has a tendency to dry out dur- ing prolonged storage or incubation, unless it is kept in plastic bags that retard the loss of moisture. (2) Exposure of the completed medium to daylight causes the release of formaldehyde gas in concentrations sufficient to inhibit growth of mycobacteria (15). (3) If 0.1% L-aspartic acid is added to enhance the niacin pro- duction of M. tuberculosis grown on agar medium, the activity of certain drugs is altered (6). (4) The preparation of this medium requires great care, espe- cially when aseptically adding the enrichment. Most media used in the mycobacteriology laboratory are commer- cially available as complete ready-to-use media, or as powdered bases that require minimal preparation and occasional addition of supple- ments. If commercial media are used, each lot should be checked with a strain of M. tuberculosis having known growth characteristics (e.g., H37Ra). The following variables contribute to the preparation of a good culture medium: purity of chemical components, care taken while preparing and sterilizing the medium and glassware, final pH, exposure of final product to excessive heat or sunlight, method and length of storage (7,8,15,20,22). 2. Preparation of Egg-Base Medium The most commonly used egg-base media are modified Lowenstein- Jensen (L-J) (5,8) and American Trudeau Society (ATS) (21). The ATS medium, which contains no asparagine or citrate, has the simplest formulation of all the egg media (10). Various selective media that contain antibiotics include L-J-Gruft (4) that contains penicillin and nalidixic acid; and L-J-Mycobactosel (17), with cycloheximide, linco- mycin, and nalidixic acid. Antimicrobial-containing isolation medium is often used in an attempt to control excessive culture contamination. This precaution is an expensive alternative and should be taken only 48 after careful evaluation of the digestion-decontamination procedure. Even if antimicrobial-containing medium is added to the isolation media used, this should be done only if controlled by the inclusion of the same basic medium without antimicrobial agents added. Although most of these media are commercially available, some laboratories prefer to make their culture media from basic ingredients. eo Modified Lowenstein-Jensen Egg Medium (5) Fresh eggs, not more than 1 week old, are cleaned by scrubbing with a hand brush in a soap solution. Let the eggs soak for 30 minutes in the soap solution. Rinse eggs thoroughly in running water and soak them for 15 minutes in 70% ethanol. Wash and scrub hands well before breaking the eggs into a sterile flask. Shake the flask by hand to homogenize the eggs. Filter eggs through four layers of sterile gauze into a sterile, graduated cylinder. (1) For salt solution, dissolve in order: Monopotassium phosphate (anhydrous) ......... 24g Magnesium sulfate -7H,0 .. .................. 0.24 g Magnesium citrate ................... ....... 06g ASParagine ......sounsssssmunss os mnvameyss as 36g Glycerol (reagent grade) ................... 12.0 ml Distilled water .......................... 600.0 ml (2) Add potato flour .................... ...... 300g (3) Autoclave at 121°C for 30 minutes. (4) Cool to room temperature. (5) Add malachite green (2% aqueous solution, freshly prepared) ..... 20.0 ml (6) Add homogenized wholeeggs ............ 1000.0 ml (7) Mix and pour into a sterile aspirator bottle or funnel with a bell attachment (test tube filling device) and dispense. (8) Place approximately 6 to 8 ml of medium into each 20 x 150-mm sterile screwcap test tube. (9) Slant tubes and coagulate by inspissation at 85°C for 50 minutes. (10) Incubate at 37°C for 48 hours as a sterility check. Medium may be stored in the refrigerator for several months if caps are tightly closed to prevent evaporation. 3. Preparation of Agar-Base Medium The most popular agar-base media are Dubos oleic acid-albumin (18), Middlebrook 7H-10 (12,13), and Middlebrook 7H-11 (3). 7H-11, a 7H-10 agar enriched by the addition of enzymatic digest of casein, stimulates more profuse growth (particularly of drug-resistant tuber- cle bacilli), but care should be taken when using 7H-11 for drug sus- ceptibility testing because components of the digested casein addi- 49 tive may affect the minimal inhibitory concentration of some anti- tuberculosis drugs (11). Mitchison et al. (16) proposed the use of selective 7H-10 or 7H-11 agars to recover tubercle bacilli from sputum without the need to decontaminate the digested sputum. McClatchy et al. (11) noted that these media were inhibitory to some species of mycobacteria and suggested that the addition of 50 pg of carbenicillin per milliliter rather than 100 pg/ml resulted in an increase in the numbers of some mycobacterial species reported. e 7H-10 Agar Medium (12) Middlebrook 7H-10 agar medium may be prepared by using com- mercially available 7H-10 agar-powdered base and Middlebrook OADC (oleic acid-albumin-dextrose-catalase) enrichment. It is best to pre- pare the medium in small quantities of 200 to 400 ml to minimize the amount of heat needed to melt the agar. Boiling the basal medium before autoclaving (either to solubilize the agar or to provide stocks of prepared base that may be stored and boiled for later use) should be avoided because the double heating produces a medium of infe- rior quality. To prepare 200 ml of complete medium, follow these directions precisely: (1) Suspend 3.6 g of Middlebrook 7H-10 basal medium in 180 ml of freshly distilled water and add 1.0 ml of reagent grade glycerol. (2) Swirl base into suspension and sterilize in the autoclave for 10 minutes at 121°C. (3) Remove medium from autoclave as soon as the pressure will allow and place in a water bath at 50 to 56°C. The medium should be clear and transparent. (4) As soon as cooled to 50 to 56°C, add 20 ml of OADC enrich- ment that has been warmed to room temperature. Do not temper the OADC in a 56°C water bath. (5) For drug media, cool to 50 to 52°C and add the required amount of drug. (6) Dispense within 1 hour after autoclaving to prevent a pre- cipitate from forming. (7) Allow to solidify at room temperature without exposure to daylight (fluorescent light is all right). Caution: Never autoclave 7H-10 medium base and store it in the refrigerator for future use. The heat required to re-melt it produces a medium of poor quality. eo 7H-11 Agar Medium (3) To prepare 200 ml complete medium: (1) Suspend 3.78 g of 7H-11 agar basal medium in 180 ml of distilled water containing 1 ml of glycerol. 50 (2) Autoclave for 15 minutes at 121°C. (3) Cool to 50 to 52°C in a water bath, add 20 ml of OADC enrichment (warmed to room temperature), and pour medium into plates. (4) For drug media, cool to 50 to 52°C; add drugs as needed, then OADC enrichment; and pour into plates. (5) Dispense within 1 hour after autoclaving. Note: Observe same precaution as for 7H-10 medium. e Mitchison’s Selective 7H-10 or 7H-11 Medium (16) Either 7H-10 or 7H-11 medium may be made “‘selective’’ (suppres- sive for most contaminants) by the aseptic addition to tempered (nonsolidified) complete medium of sterile solutions of the following antimicrobial agents in the final concentrations listed: Agent Final Concentration Polymyxin B 200 units/ml Amphotericin B 10 pg/ml Carbenicillin 50 pg/ml Trimethoprim 10 or 20 pg/ml 4. Preparation of Middlebrook 7H-9 Liquid Medium (12) Liquid medium is used almost daily in the mycobacteriology labora- tory for subculturing stock strains, picking single colonies, and prepar- ing inoculum for drug susceptibility tests and other in vitro tests. Middlebrook 7H-9 medium may be prepared from commercially avail- able powdered base that is supplemented with Middlebrook ADC (albumin-dextrose-catalase) enrichment after sterilization of the basal liquid medium. Glycerol or Tween 80 may also be added, but not both together. Glycerol may be included in the basal medium (0.2% final concentration) as a carbon source. Mycobacteria have a ten- dency to clump in liquid medium; Tween 80, a wetting agent that encourages more homogeneous growth, may be included in the basal medium at a final concentration of 0.05%. Uniform, homogenous suspensions of mycobacteria are important in preparing dilutions for drug-susceptibility tests and in vitro tests. To prepare 1000 ml of complete medium: (1) Suspend 4.7 g of the dehydrated medium in 900 ml of distilled water containing 2 ml of glycerol, or 0.5 g (ml) of Tween 80. Do not use Tween 80 and glycerol together. (2) Autoclave at 121°C for 15 minutes. (3) Remove from autoclave and cool to 45°C (use a water bath). (4) Add aseptically 100 ml of ADC enrichment to the cool (45°C) basal medium. (5) Dispense aseptically 5-ml amounts into 20 x 150-mm screwcap test tubes. 51 (6) The completed medium stores almost indefinitely if caps are tightened so the liquid does not evaporate. 5. Storage of Media (1) Protect any medium from direct light at all times. (2) Place plates in clean plastic bags or sterile dry containers. (3) Tighten caps of screwcap tubes before storing. (4) Refrigerate any medium kept for longer than 1 week to minimize dehydration. (5) Discard any media that show signs of contamination or dehydration. D. Guidelines for Optimal Culture Growth When attempting to culture mycobacteria from digested-decon- taminated clinical specimens, remember that: « Different species of mycobacteria, as well as different strains of the same species, may exhibit different growth require- ments Mycobacteria may be clumped and unevenly distributed in clinical specimens « The digestant-decontaminant often has harmful effects on mycobacteria e The specimen itself, or something in it, may be toxic for the mycobacteria 1. Preparation of Media Use reagent grade chemicals and ingredients (unless otherwise specified), and follow directions carefully. Do not overheat any medium during preparation. Carefully monitor temperatures of autoclaves and inspissators. Do not expose completed medium to direct daylight during or after preparation; this is especially true of 7H-10 and 7H-11 agar-base media. Do not skimp on the quantity of medium placed in containers; use 8 ml of egg medium in 20 x 150-mm screwcap test tubes and 20 ml of agar medium in 15 x 100-mm dishes. 2. Inoculation of Media It is recommended that digested-decontaminated specimens be inoculated, both undiluted and after dilution (107'), onto multiple units of at least two different kinds of basal medium. The tenfold dilution acts to reduce the concentration of any possible toxic material(s) in the specimen. Inoculate at least four units of an egg- base medium and one biplate of an agar-base medium. 52 The sediment from the processed specimen is spread evenly over the entire surface of the tubed medium using a bacteriologic loop, a disposable applicator stick, a capillary pipette, or a serologic pipette. Plated medium is often inoculated with a capillary or serologic pipette by touching three distinct (equally spaced) drops of inoculum to the medium just before they “free fall’”’ from the pipette. Plates and tubes of inoculated media may be left at room tempera- ture for several hours until the fluid inoculum dries or is absorbed. Then, seal plates in CO,-permeable polyethylene plastic bags to mini- mize drying and keep them shielded from daylight to avoid the for- mation of toxic formaldehyde (15). 3. Incubation of Cultures e Tubed media should be incubated in a slanted position with screwcaps loose for at least 1 week to ensure even distribution of inoculum. Thereafter, if space is needed, tubes may be placed upright with the caps tightened to minimize evaporation and drying of media. e Plated medium, placed in CO,-permeable plastic bags, should be incubated with the medium-side down if all the inoculum has not been absorbed. Do not stack plates more than six high. e All media should be examined within 5 to 7 days after incubation to permit early detection of rapidly growing mycobacteria and to enable prompt removal of contaminated cultures. Thereafter, all cul- tures should be incubated for 8 weeks with weekly examination for evidence of growth. Prolonged incubation (10 to 12 weeks or more) may be necessary in selected cases, but usually this may be avoided by processing more specimens from the same patient. 4. Atmosphere for Incubation All media inoculated for primary isolation should be incubated in an atmosphere of 10% CO, and 90% air (1). The various Middlebrook agars require a CO, atmosphere to ensure growth. Although CO, is not essential to initiate growth on egg-base medium, it does stimu- late earlier and more luxuriant growth (4, 19). The CO, concentration in the incubator should be monitored daily with a Fyrite CO, gas analyzer or some other method for determining CO, concentration. Often a separate CO, incubator is not available for one of the spe- cial temperatures (25 to 33°C or 40 to 42°C), or only a small amount of CO, incubation space is needed. In either case, one of the following alternative methods may be employed: (1) Attach inlet and outlet petcocks to an airtight metal or a plastic box built to fit on the shelf of an incubator main- tained at the desired temperature. The box that contains 53 (2) the incubating cultures, should be flushed daily with a com- pressed mixture of 10% CO, and 90% air. A CO,-impermeable Mylar plastic bag (7x14 in.) may be “converted’’ to a CO, incubator as follows (2, 7): Affix a 1 x 1-inch square of 1/16-inch thick neoprene rubber to the Mylar bag (old automobile innertube is also accept- able) using a 2 1/2-inch square of silver duct tape. Place 6 to 8 plates in the bag; twist and seal the opening with mask- ing tape. Construct a vacuum line using a 1 liter sidearm Erlenmeyer flask containing disinfectant, rubber pressure tubing, and a 20-gauge needle attached to the barrel of a 1-ml plastic syringe, as shown in figure 14. Attach the unit to a vacuum line and carefully insert the needle through the rubber gasket to evacuate air from the Mylar bag. Air from the bag should pass through the disinfectant reser- voir of 2% Amphyl containing a few milliliters of an antifoam solution. Draw a vacuum until the plastic bag wraps close around the plates. Do not leave vacuum unattended because plates may be crushed and broken by excessive vacuum. After air is evacuated from the bag, remove the needle and reinflate the bag with a mixture of 10% CO, and 90% air. This may be done by using another 20-gauge needle on a syringe barrel attached with rubber tubing to a compressed gas cylinder of 10% CO, and 90% air. The cylinder of com- pressed gas must be fitted with a step-down pressure gauge to enable pressure control of 1 or 2 psi. Because metaboliz- ing mycobacteria rapidly deplete the CO,, the Mylar bag incubator must have the gas contents exchanged 3 times a week to maintain the required CO, concentration. FIGURE 14. Mylar bag CO, incubator 54 5. Temperature of Incubator The optimum temperature for incubating media inoculated with most specimens from the human body is 35 to 37°C. Exceptions are made for specimens obtained from skin or superficial lesions sus- pected to contain M. marium, M. ulcerans, or M haemophilum. These organisms multiply best at 25 to 33°C. Cultures suspected to contain M. avium or M. xenopi exhibit optimum growth at 40 to 42°C. 6. Precautions To obtain maximum growth on culture media, it is imperative that the decontamination procedure be followed precisely. Inefficient cen- trifugation or overexposure to the digestion-decontaminating agent may lead to considerable reduction in the numbers of mycobacteria that were present in the submitted specimens (9). To minimize the accumulation of moisture inside plates of inocu- lated media during incubation: (1) Let the inoculum evaporate or be absorbed into the medium before sealing plates in individual plastic bags. (2) Do not inoculate a plate of medium with a wet surface. The day before plates are to be inoculated, remove them from the refrigerator and allow them to warm to room tempera- ture. (3) Use styrofoam sheets (1/2-inch thick) or corrugated card- board to insulate plastic plates from metal shelves in the incubator. Plates in direct contact with metal shelves seem to accumulate more moisture than those insulated from the metal shelving. (4) Avoid incubating plates on shelves adjacent to incubator door. Fluctuation in temperature of air currents created by frequent opening of door may cause condensation within plates or plastic bags. (5) When reading groups of plates stacked in large plastic bags, open the bags immediately after removal from the incuba- tor. This will minimize condensation within the bag. (6) If condensate forms on the lids of plates during micro- scopic examination, use a warming pad (e.g., electric heat- ing pad) to warm plate lids so that condensate is reduced. 55 REFERENCES 1. 15. 16. 17. 18. 19. 20. 21. 22. Beam RE, Kubica GP. Stimulatory effects of carbon dioxide on the primary isolation of tubercle bacilli on agar containing medium. Am J Clin Pathol 1968;50:395-7. . Cohn ML, Middlebrook G. Carbon dioxide-impermeable plastic bags for artificial cultivation of tubercle bacilli. Am Rev Respir Dis 1963;87:292. . Cohn ML, Waggoner RF, McClatchy JK. The 7H-11 medium for the culture of mycobacteria. Am Rev Respir Dis 1968;98:295-6. . Gruft H. Isolation of acid-fast bacilli from contaminated specimens. Health Lab Sci 1971;8:79-82. . Holm J, Lester V. Diagnostic demonstration of tubercle bacilli. Public Health Rep 1947;62:847-54. . Kilburn JO, Stottmeier KD, Kubica GP. Aspartic acid as a precursor for niacin synthesis by tubercle bacilli grown on 7H-10 agar medium. Am J Clin Pathol 1968;50:582-6. . Kubica GP, David HL. The Mycobacteria. In: Sonnenwirth AC, Jarett L, eds. Gradwohl’s clinical laboratory methods and diagnosis. 8th ed. St. Louis: C.V. Mosby Co., 1980:1693-730. . Kubica GP, Good RC. The genus Mycobacterium (except M. leprae). In: Starr HP, Stolp H, Truper HG, Balows A, Schlegel HG, eds: The prokaryotes: a handbook on habitats, isolation and identification of bacteria. New York: Springer-Verlag, 1981:1962-84. . Kubica G, Kent PT. The sputum digestion process in the mycobacteriology laboratory: contributions of centrifugal efficiency and digestant toxicity. (In preparation, 1985). . Marks J, Thomas CH. Notes on the cultivation of tubercle bacilli. The monthly Bulletin of the Ministry of Health and the Public Health Lab Service 1958;17:194-202. . McClatchy JK, Waggoner RF, Kanes W, Arnick MS, Bolton TL. Isolation of myco- bacteria from clinical specimens by use of a selective 7H-11 medium. Am J Clin Pathol 1976;65:412-5. . Middlebrook G, Cohn ML. Bacteriology of tuberculosis: laboratory methods. Am J Public Health 1958;48:844-53. . Middlebrook G, Cohn ML, Dye WE, Russell WF Jr, Levy D. Microbiologic proce- dures of value in tuberculosis. Acta Tuberc Scand 1960;38:66-81. . Middlebrook G, Cohn ML, Shaefer WB. Studies on isoniazid and tubercle bacilli. Ill. The isolation, drug-susceptibility and catalase testing of tubercle bacilli from isoniazid-treated patients. Am Rev Tuberc 1954;70:852-72. Miliner RA, Stottmeier KD, Kubica GP. Formaldehyde: a photo-thermal activated toxic substance produced in Middlebrook 7H-10 medium. Am Rev Respir Dis 1969;99-603-7. Mitchison DA, Allen BW, Carol L, Dickinson JM, Aber VR. A selective oleic acid albumin agar medium for tubercle bacilli. J Med Microbiol 1972;5:165-75. Petran El, Vera HD. Media for selective isolation of mycobacteria. Health Lab Sci 1971,8:225-30. Runyon EH, Karlson AG, Kubica GP, et al. Mycobacterium. In: Lennette EH, Spaulding EH, Truant JP, eds. Manual of clinical microbiology. 2nd ed. Washington: American Society of Microbiology, 1974. Whitcomb FC, Foster MC, Dukes CN. Increased carbon dioxide tension and the primary isolation of mycobacteria. Am Rev of Respir Dis 1962;86:584-6. Willis HS, Cummings MM. Diagnostic and experimental methods in tuberculosis. 2nd ed. Springfield, Ill: Charles C Thomas, 1952. Woodruff CE, Crombie D, Wooley JG, Medlar E, Steenken W. American Trudeau Society: Committee of evaluation of laboratory procedures. Am Rev Tuberc 1946;54:428-32. Youmans GP. Tuberculosis. Philadelphia: W.B. Saunders Co., 1979. 56 E. Acid-Fast Microscopy Microscopy is perhaps the easiest and most rapid procedure that can be performed in the laboratory to detect the presence of acid-fast bacilli. It is, however, much less sensitive than culture for detecting mycobacteria. David (9) has estimated there must be 5,000 to 10,000 bacilli per milliliter of sputum to permit their routine detection in stained smears. Earlier quantitative studies that compared smear with culture results provided comparable estimates (14, 25). In contrast to microscopy, culture techniques have been estimated to detect 10 to 100 viable mycobacteria per milliliter of sample. In spite of this quantitative discrepancy in sensitivity, examination of stained smears of sputum, or other clinical material, can be helpful in several ways: (a) it provides a presumptive diagnosis of mycobacte- rial disease: (b) it enables the rapid identification of most infectious cases, i.e., those smear positive (11, 18, 20); (c) it may be used to follow the progress of tuberculous patients on chemotherapy; (d) it is of vital importance in regard to the patient's discharge from the hospital, or return to gainful employment (7); (e) it can confirm that cultures growing on media are indeed acid-fast; and (f) it is useful in determining appropriate dilutions of sediments for direct drug sus- ceptibility tests. 1. General Comments and Precautions Microscope slides used for acid-fast staining should be new, clean, and unscratched. Acid-fast material from previous smears may be retained on old, washed, and reused slides and, thus, may lead to false-positive reports. One end of the slide should be labeled with the patient's name and/or hospital identification number and date. The specimen to be smeared should be spread over an area approxi- mately 1 x 2 cm, as illustrated in figure 15. FIGURE 15. Smear area To smear the specimen over a larger area would be self-defeating. If a microscopist examines 100 to 300 fields on a stained smear 1 x 2 cm, only 1/100 to 4/100 of the entire area of the smear is observed (i.e., 1% to 4% of the smear area), depending on the magnification used for scanning (9, 21). A larger smear, examined in the same manner, would only further reduce examining efficiency. Keep the area of the smear fairly small; 1 x 2 cm is satisfactory. 57 Use a bacteriological loop, applicator stick, or pipette to make smears. The loop may be reused after it is dipped into an alcohol sand flask and sterilized in the flame of a Bunsen burner, but applica- tor sticks and pipettes should be discarded after each smear is prepared. Let the smear air dry and then heat-fix it. Use either an electric slide warmer at 65 to 75°C for at least 2 hours (overnight is acceptable), or pass the slide 3 to 4 times through the blue cone of a burner flame as for other bacteriological smears. Heat-fixing does not always kill the mycobacteria, so handle smears carefully. Discard stained slides into covered pans containing disinfec- tant solution and autoclave them before disposal, or discard them into 3% amphyl solution overnight. Many acid-fast staining proce- dures have been described, including hot and cold staining methods and those which employ fluorescing dyes. All are based on the prop- erty of mycobacteria to retain the primary stain, even after exposure to strong mineral acids or acid-alcohol—thus the term ‘acid-fast bacilli.” Prepare and stain smears carefully to avoid such problems as poorly stained preparations and false-positive or false-negative reports. The following suggestions may be helpful in avoiding some of these problems. (1) Avoid insufficient destaining with acid-alcohol. e Most mycobacteria are strongly acid-fast and are not easily decolorized e Smears that are too thick are difficult to destain, and the subsequent counter stain may obscure acid-fast bacilli (2) Use a contrasting counterstain. e Counterstain should not be so intense that it “‘hides”’ the stained mycobacteria e Color-blind individuals may find fluorochrome-stained smears easier to examine (3) Be aware of possible sources (environmental or procedural) of false-positive smears. Common sources of acid-fast bacilli that contribute to this problem are: e Tap water and infrequently cleaned distilled water reservoirs. Check these sources from time to time to see if acid-fast bacilli can be detected. Tap water may be especially problematic after heavy rainstorms. The distilled water reservior that is always left on the shelf and refilled from an outside supply source is espe- cially suspect e lce-making machines. Ice is often used to facilitate passage of gastric tubes; if the tap water used to make the ice is contaminated with mycobacteria, it could lead to false-positive smears or cultures 58 « Transfer of material from slide to slide in bulk-staining tanks. To avoid this problem, stain slides individually « Transfer of ““positive’’ flakes from thick slides to other slides via immersion oil. Do not make thick smears; carefully heat-fix smears; wipe oil immersion lens between slides; allow oil to free-fall from applicator onto the smear instead of touching applicator to the slide 2. Smear Preparation The method of smear preparation most used in developing coun- tries and in those laboratories where a rapid smear examination is required is the direct smear. Select the cheesy, necrotic, blood-tinged particles in the specimen because they are the most likely to produce postive smear results. There are several ways to prepare concentrated smears and some of these are discussed in the section on “Isolation Procedures.” If the specimen is also to be used for cultural identification, the sediment from the digestion-decontamination procedure is first inoculated onto culture media before being smeared onto a slide. If only smears are to be made, as in the Level | laboratory, the risk of infecting labora- tory personnel is greatly reduced by using a hypochlorite concentra- tion method. Clorox or other 5% to 6% hypochlorite solution is bacte- ricidal for most organisms in the specimen including mycobacteria; therefore, after being treated with Clorox, the specimen may be han- dled safely without a safety cabinet. Clorox may cause disintegration of the acid-fast bacilli if it is allowed to act longer than 15 minutes; therefore, smears should be prepared, stained, and examined promptly. 3. Sodium Hypochlorite Method for Concentration of Mycobacteria (6, 8 19, 21) e Materials Clorox or other commercial household bleach containing 5% to 6% sodium hypochlorite (NaOCl). Screwcap plastic centrifuge tubes, preferably 50 ml. Centrifuge and head to provide 3000 x g. e Procedure (1) Mix equal volumes of sputum and Clorox (5% sodium hypochlorite) in a screwcap centrifuge tube. (2) Tighten cap and shake the tube to liquefy the specimen. A test tube mixer may be used. (3) Let the mixture stand for 15 minutes at room temperature. 59 (4) Add sterile water to the 50-m| mark on the tube. (5) Centrifuge the tube(s) at 3000 x g for 15 minutes. (6) Decant the supernatant fluid and retain the sediment. (7) Suspend the sediment in several drops of water and pre- pare the smear by spreading a drop of the resuspended material over an area 1 x 2 cm. Allow to air dry. (8) Fix the smear by passing the slide through the blue cone of a Bunsen burner flame about three times. (9) Perform acid-fast staining procedure as described below. Another method for concentrating virtually all of the AFB in a spu- tum into one small area is the recently described polycarbonate mem- brane filter technique (22a). Using a vacuum pump, a small volume of sodium hypochlorite-digested sputum is deposited ontothe membrane filter; the filter is next inverted, fixed onto a microscope slide, and stained in place. Before the final rinsing step, the membrane filter is carefully lifted away from the now-adhered smear. Consult the origi- nal reference for precise details. 4. Staining Methods Acid-fast staining procedures are of two general types: those using basic fuchsin dyes and those using fluorescing dyes. Some of the commonly used staining methods are outlined below. e Basic Fuchsin Acid-Fast Stains (1) Ziehl-Neelsen is a HOT acid-fast stain (5, 9, 21) MATERIALS: Basic fuchsin Ethanol, 95% Phenol crystals Hydrochloric acid, concentrated Methylene blue chloride Water, distilled PREPARATIONS: (a) Fuchsin—Dissolve 0.3 g of basic fuchsin in 10 ml of 95% ethanol. (b) Phenol—Dissolve 5.0 g of phenol crystals in 100 ml of water (gentle heat may be needed). (c) Carbol fuchsin—Mix solution (a) with 90 ml of solution (b). (d) Acid alcohol—Carefully add 3 ml of concentrated hy- drochloric acid to 97 ml of 95% ethanol; mix gently. (e) Methylene blue—Dissolve 0.3 g of methylene blue chloride in 100 ml of distilled water. PROCEDURE: (a) Prepare smear as described; allow to air dry. 60 (2) (b) (c) (d) (e) (f) (g) (h) (i) (j) (k) Heat-fix smear either on an electric slide warmer at 65 to 75°C for at least 2 hours or pass slide through Bun- sen burner flame as for other bacteriological smears. DO NOT OVERHEAT. Cover smear with a 2 x 3-cm piece of filter paper to hold the stain on the slide and to minimize the precipi- tation of dye crystals onto the smear. Flood the paper strip with carbol fuchsin. Heat the slide to steaming with a Bunsen burner or an electric staining rack; let stand 5 minutes. If the smear dries, add more stain but do not reheat. Use forceps to remove paper strips from slides and to place them in discard container. Wash slides with water (use tap water or water from reservoir bottles). Flood smear with acid-alcohol; allow to destain for 2 minutes. Wash smear again with water and drain. Flood slide with methylene blue and counterstain for 1 to 2 minutes. Rinse with water, drain, and air dry. DO NOT BLOT. Examine smear under oil immersion objective lens of the microscope (ca. 1000 X). Kinyoun is a cold acid-fast stain (9, 15, 17, 21). MATERIALS: Basic fuchsin Ethanol, 95% Phenol crystals Hydrochloric acid, concentrated Methylene blue Water, distilled PREPARATIONS: (a) Basic fuchsin—Dissolve 4 g of basic fuchsin in 20 ml of 95% of ethanol. (b) Phenol—Dissolve 8 g phenol crystals in 100 ml of dis- tilled water. Heat may be required to effect solution. (c) Carbol fuchsin—Mix solutions (a) and (b). (d) Acid alcohol—Carefully add 3 ml of concentrated hy- drochloric acid to 97 ml of 95% ethanol; mix gently. (e) Methylene blue—Dissolve 0.3 g of methylene blue chloride in of 100 ml of distilled water. PROCEDURE: (a) Prepare smear as described and allow to air dry. 61 (b) Heat-fix smear on an electric slide warmer at 65 to 75°C for at least 2 hours or pass slide through flame of Bunsen burner as for other bacteriological smears. DO NOT OVERHEAT. (c) Cover smear with a 2 x 3-cm piece of filter paper to hold the stain on the slide and to filter out any undis- solved crystals of dye. (d) Flood the paper strip with Kinyoun carbol fuchsin and stain for 5 minutes. DO NOT HEAT. Heat is not neces- sary because of the increased concenteration of both the primary stain and the phenol. (e) Use forceps to remove filter paper strips from slides and to place them in discard container. Rinse slide with tap water or bottled water and drain. (f) Destain smear with acid alcohol for 2 minutes. (g) Rinse again with water and drain. (h) Flood smear with methylene blue and counterstain for 1 to 2 minutes. (i) Rinse, drain, and air dry. DO NOT BLOT. (j) Examine smear under oil immersion objective lens of the microscope (ca. 1000 X). e Fluorochrome Acid-Fast Stains There is a trend toward greater use of fluorescent acid-fast staining procedures because smears so stained may be examined more rap- idly and results ‘appear’ to be more sensitive than those obtained with the basic fuchsin procedures. Fluorochrome-stained smears are generally scanned at 250 X to 450 X while fuchsin-stained smears are examined at 800 X to 1000 X. The difference in magnification alone provides an obvious savings of time for the microscopist. The lower magnification used to scan fluorochrome-stained smears enables the viewer to observe a much larger area (4 to 10 times) of the smear during the same time period than could be achieved with fuchsin- stained preparations examined at a higher magnification; this contrib- utes to the illusion of greater sensitivity for the fluorescent acid-fast stains. In addition, it is easier for the eye to detect a yellow bacillus fluorescing against a dark (potassium permanganate) or dark red (acridine orange) background, rather than a red, fuchsin-stained bacil- lus in the middle of surrounding methylene blue-counterstained tis- sue debris. Fluorochrome-stained smears should be observed within 24 hours of staining because of the possibility of fading of the fluorescence (21). Stained smears may be stored at 5°C overnight to minimize the loss of fluorescence, but if at all possible, do not stain smears unless they can be observed within 24 hours. 62 (1) Auramine 0 Fluorescence Acid-fast Stain (4, 17, 21) MATERIALS: Auramine 0 Phenol crystals Hydrochloric acid, concentrated Ethanol, 70%, 95% Potassium permanganate Water, distilled PREPARATIONS: (a) (b) (c) (d) (e) Auramine 0 - Dissolve 0.1 g of auramine 0 in 10 ml of 95% ethanol. Phenol - Dissolve 3.0 g of phenol crystals in 87 ml of distilled water. Mix solutions (a) and (b). Acid alcohol - Carefully add 0.5 ml of concentrated hydrochloric acid to 100 ml of 70% ethanol. Potassium permanganate - Dissolve 0.5 g of potas- sium permanganate (KMnO,) in 100 ml of distilled water. PROCEDURE: (a) (b) (c) (d) (e) (f) (9) Prepare smear as described earlier and allow to air dry. Fix smear on an electric slide warmer at 65 to 75°C for at least 2 hours, or use a Bunsen burner flame, as for other bacteriological smears. DO NOT OVERHEAT. Flood smear with auramine 0 solution and allow to stain for 15 minutes, making certain that the staining solution remains on the smear. Do not apply heat to smear. Do not use filter paper strips. Rinse smear with chlorine-free water and drain. Chlo- rine may interfere with fluorescence; therefore, rinse with distilled or deionized water. Flood smear with acid alcohol and allow to destain for 2 minutes. Rinse again and drain smear. Flood smear with potassium permanganate* and counterstain for 2 minutes. Time is critical with potas- sium permanganate because counterstaining for a longer time may quench the fluorescence of the acid- fast bacilli. *Acridine orange may be used in place of potassium permanganate as the counterstain (22): Dissolve 0.01 g of anhydrous dibasic sodium phosphate (Na,HPO,) in 100 ml of distilled water; add and dissolve 0.01 g of acridine orange. Counterstaining time remains 2 minutes. 63 (h) Rinse, drain, and allow smear to air dry. (i) Examine smear with fluorescence microscope as soon as possible. 5. Examination of Smears Careful smear examination is an essential part of today’s tuberculo- sis control program. The microscopist’s training should emphasize the importance the clinician attaches to smear results in deciding to discharge a patient to outpatient treatment or to gainful employment (3,7, 13). To attain excellence in microscopic examination, one should have a good microscope and a comfortable work area. Methods of reading smears vary from laboratory to laboratory. At CDC, no time limit has been set for examining a smear. Instead, a system has been adopted that ensures that a representative area of the smear is examined. To ensure that an area is covered only once, the smear should be searched in an orderly manner by making a series of three parallel sweeps the length of the smear or nine parallel sweeps the width of the smear (see figure 16). Each field should be searched thoroughly, with a rapid change to the next field. These procedures should be followed regardless of how much time it takes. In this manner, the microscopist should see 60 to 100 fields in one sweep, or as many as 300 fields if it is necessary to examine the entire three or nine parallel sweeps of the smear. If the smear is moderately or heavily positive, then fewer fields need be examined, and a report of “positive’’ may be made even though the entire smear has not been examined. | == | EY FIGURE 16. Recommended method for examining acid-fast stained smears. 64 Acid-fast bacilli in specimens usually are rod-shaped, 1- to 10-um long and 0.2- to 0.6-um wide; but they also may appear coccoid or filamentous (long, slender and even branching). They are frequently bent and may contain heavily stained areas called beads, or alternat- ing stained and clear areas that make them appear banded. See color plates 1 and 2. From pure cultures, individual rods of M. tuberculosis may be aggregated side by side and end to end to form “cords” (color plate 3). This is often the case in smears made from the turbid fluid at the bottom of a Lowenstein-Jensen slant on which M. tuberculosis is growing. As smears of pure cultures reveal, not all of the mycobacteria in a given smear are necessarily acid fast, so careful attention must be paid to cellular morphology and size before labeling a smear, “mixed or contaminated.” Then, too, some species of the genera Norcadia and Corynebacterium, and even some bacterial and fungal spores, may be acid fast (2). In fluorescence acic-fast microscopy, tissue or medium debris may fluoresce and be mistaken for acid-fast bacilli. By examining smears frequently and becoming familiar with the mor- phology of acid-fast organisms, the microscopist learns to distin- guish acid-fast bacilli from these other, sometimes confusing, organ- isms and artifacts. COLOR PLATE 1. Branching filaments with beading and small coccoid forms. (21) COLOR PLATE 2. Alternating clear and pink-stained parts of the bacilli that show the characteristic of banding. Note the dark-stained parts called beads. COLOR PLATE 3. Strands of bacilli in cords. (19) 65 General information on the use and maintenance of equipment for light microscopy and fluorescence microscopy may be found in two CDC manuals (21, 23). More specific and detailed information is avail- able from the microscope manufacturer or supplier. 6. Reporting Smear Results Smear results must be quantified to be meaningful. The following method provides semiquantitative reporting in roughly tenfold incre- ments for fuchsin-stained smears observed at 1000 X (1, 9, 21). Number of AFB seen Report None No AFB observed 1-2/300 fields + % 1-9/100 fields 1+ 1-9/10 fields 2+ 1-9/field 3+ >9/field 4+ When fluorochrome staining methods are used, smears are exam- ined at much lower magnifications than those commonly used for fuchsin-stained smears. Because of this, each field examined under fluorescence microscopy (examination at 250 X to 630 X) has a larger area than that seen with bright field microscopy (ca. 1000 X). Use of different staining techniques or, more particularly, different total magnifications for examination of stained smears, can be confusing to the attending clinician who may use changes in the quantitative smear report as a measure of patient response to treatment. In other words, a report based upon a fluorochrome-stained smear examined at 250 X may contain much larger numbers of bacilli than a similar report from the same specimen stained with carbol fuchsin and exam- ined at 1000 X. To minimize confusion that conceivably could occur when mixed modes of staining and variable magnifications are used for smear examination and quantitative reporting of results (i.e., = to 4+, as shown above), Smithwick (21) suggested that any number of AFB observed under fluorochrome stain could be divided by a “magnification-factor’’ number to yield an approximate number that might be observed if the same smear were examined under 1000 X after carbol fuchsin stain; i.e., the ratings of + to 4+ could be equated among the various magnifications used for smear examination in the mycobacteriology laboratory. To avoid making these computations, table 2 indicates the approximate number of fields that must be observed under fluorescent stains (examined from 250 X to 630 X) to *Any number of AFB less then 3/300 fields is doubtful; either reexamine the smear or make a new smear from the same or another specimen (if available). 66 make them equivalent to 300 fields seen under fuchsin stains exam- ined at 1000 X. Also listed are the comparable numbers of organisms per field(s) needed to equate the fluorochrome smears to the scores of + to 4+ reported under fuchsin stain. For purposes of comparison, the Smithwick ““magnification-factors’’ also are listed. By using table 2, the final report should be comparable from person to person or laboratory to laboratory regardless of the stain or magnification used for examination. A similar technique already is used by Grosset and Truffot-Pernot (12). On the final report: e State the staining method used and the number of acid-fast bacilli seen on the smear. « Observe only enough fields to get an average number of acid-fast organisms. e Count as one a clump of bacilli that are touching. Give separate counts to separate organisms. A clump of bacilli is considered a single colony-forming unit; this is important to know when deter- mining dilutions of sediment for direct drug susceptibility tests. « Note any large numbers of clumps on the report, thus indicating that the actual number of acid-fast organisms is larger than that reported. « REPORT ONLY THE NUMBER OF ACID-FAST BACILLI SEEN: DO NOT TRY TO ““SPECIATE"” WITH MICROSCOPY ALONE. e Counts of less than 3 per 300 fields by fuchsin stains (or 3 per varying numbers of fields by fluorochrome stain; see table 2) are not considered positive. A report of “‘doubtful” may be sent or, if TABLE 2. Smear evaluation oo | Number of AFB Observed by Fluorochrome at Magnification ho 20x ~ 450X ~ 630X Report 1000X K/K** RWS** K/K RWS K/K RWS - TT TTT [Nears 0 0 0 0 0 0 0 seen 1.2/300F 1 1230F 12m 1.2/130F Doubtful; (3 sweeps)”, (1 sweep)” Divide (1% sweeps)” Divide (2 sweeps) ® Diviiie repeat 1.9/100F 1.9/10F the 1 5.18/50F the | 5 18/100F Hig 1+ _ $b Observed 1 eeepy® Observed ji sweeps)” | Observed | 1.9/10F 1.9/F Count | 4 36/10F Count | 5 18/10F Count | 4 19F 10-90/F ay 4.36/F By 2.18/F BY 3+ >9/F >90/F > 36/F > 18/F 4+ * In all cases, one full sweep refers to scanning the full length (2 cm) of a smear 1 x 2 cm ** RWS = R.W. Smithwick; K/K = Kent/Kubica. 67 the specimen is still available, a second smear may be prepared, stained, and examined. In either case, request another specimen. If the offending Mycobacterium has already been identified, then smear examinations provide a rapid means to monitor patient response to therapy without resorting to culture. If the invasive Mycobacterium has not been identified, then the clinical specimen must be submitted for cultural identification before the acid-fast organ- isms can be specifically identified. 7. Detecting the Source of False-Positive Acid-Fast Smears (10) An increase (>2%) in the number of specimens positive by smear but negative on culture (that cannot be associated with positive response to treatment) suggests that “something” in the staining procedure (usually water) may be contaminated with acid-fast bacilli. Often, the source of contamination yields so few acid-fast organisms that they are difficult to detect without membrane filtration or high- speed centrifugation. When such equipment is not available to con- centrate the acid-fast contaminant, it may be necessary to suspend a “flocculating medium” in the water to facilitate sedimentation of the mycobacteria during centrifugation. A Candida species serves as an excellent medium because these cells lose their structural integrity and viability during digestion and decontamination, thereby posing little interference either in smear examination or in contamination of culture media. To the water (or other fluid) suspected to be contam- inated, add Candida cells to a concentration of 10° cells per milliliter. Centrifuge the suspension at 3000 X g for 15 minutes, pour off the supernatant fluid, prepare and stain smear(s) for microscopic exam- ination, and inoculate culture media with the sediment. In addition to water, all other components of the digestion and staining procedure should be examined as sources of contaminating mycobacteria; digestant solutions, buffer or water diluents, bovine albumin solution or powder, and staining reagents (pure chemicals and solutions) are examples. Do not overlook the possibility that the microscopist may be a bit overzealous in reporting positive results (this may be especially true of those who use magnifications of 250 X to scan fluorochrome smears and do not confirm suspected bacillary morphology by examination at 400 X to 630 X). If you question smear results, control positive smear reports with examination by a second, experienced reader. 68 REFERENCES 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. . American Thoracic Society. Diagnostic standards and classification of tuberculosis and other mycobacterial diseases. 14th ed. New York: American Thoracic Society, 1981. (Also available from State or local Lung Associations.) . Barksdale L, Kim KS. Mycobacterium. Bacteriol Rev 1977; 41:217-372. . Bates JH. Ambulatory treatment of tuberculosis: an idea whose time has come. Am Rev Respir Dis 1974;109:317-9. . Bennedsen J, Larsen SO. Examination for tubercle bacilli by fluorescence micros- copy. Scand J Respir Dis 1966;47:114-20. . Bishop PJ, Neuman G. The history of the Ziehl-Neelsen stain. Tubercle 1970; 51:196-206. . Cameron GM, Castles R. Detection of tubercle bacilli in sputum: application of the NaOH alum method, the clorox method, and the direct smear. J Lab Clin Med 1946;31:361-4. . Cashman HH, D’Esopo ND, Dickinson W Jr, Houk VN, Muchmore HG, Sbarbaro JA, Wolinsky E. Bacteriologic standards for the discharge of patients Am Rev Respir Dis 1970;102:470-3. . Corper HJ, Nelson CR. Methods for concentrating acid-fast bacilli. Am J Clin Pathol 1949;19:269-73. David HL. Bacteriology of the mycobacterioses. Atlanta: Centers for Disease Control, PHS, HEW, 1976. (Available from Superintendent of Documents, U.S. Government Printing Office, Washington, D.C. 20402.) Dizon D, Mihailescu C, Bae HC. Simple procedure for detection of Mycobacterium gordonae in water causing false-positive acid-fast smears. J Clin Microbiol 1976;3:211. Geiger FL, Kuemmerer JM. Tuberculosis casefinding among contacts in seven South Carolina counties. Public Health Rep 1963;78:663-8. Grosset J, Truffot-Pernot C. The role of the laboratory in the diagnosis and treat- ment of tuberculosis. Bull Int Union Tuberc 1982;57:226-34. Gunnels JJ, Bates, JH, Swindoll H. Infectivity of sputum-positive tuberculosis patients on chemotherapy. Am Rev Respir Dis 1974;109:323-30. Hobby GL, Holman AP, Iseman MD, Jones JM. Enumeration of tubercle bacilli in sputum of patients with pulmonary tuberculosis. Antimicrob Agents Chemother 1973;4:94-104. Kinyoun JJ. A note on Uhlenhuth’s method for sputum examination for tubercle bacilli. Am J Pub Health 1915;5:867-70. Kubica GP, Kent PT. The sputum digestion process in mycobacteriology: centrifu- gal efficiency and digestant toxicity. (In preparation, 1985.) Lennette EH, Spaulding EH, Truant JP. Manual of clinical microbiology. 2nd ed. Washington, D.C.: American Society for Microbiology, 1974. Loudon RG, Williamson J, Johnson JM. An analysis of 3485 tuberculosis contacts in the city of Edinburgh during 1954-55. Am Rev Tuberc 1958; 77:623-43. Oliver J, Reusser TR. Rapid method for the concentration of tubercle bacilli. Am Rev Tuberc 1942;45:450-2. Shaw JB, Wynn-Williams N. Infectivity of pulmonary tuberculosis in relation to sputum status. Am Rev Tuberc 1954;69:724-32. Smithwick RW. Laboratory manual for acid-fast microscopy. 2nd ed. Atlanta: Cen- ters for Disease Control, 1976. Smithwick RW, David HL. Acridine orange as a fluorescent counterstain with the auramine acid-fast stain. Tubercle 1971; 52:226-31. 22a.Smithwick RW, Stratigos CB. Acid-fast microscopy on polycarbonate membrane filter sputum sediments. J Clin Microbiol 1981;13:1109-13. 69 23. Strong BE, Kubica GP. Isolation and identification of Mycobacterium tuberculosis. Atlanta: Centers for Disease Control, PHS, HHS, 1981. 24. Tazir M, David HL. Evaluation of the chloride and bromide salts of cetylpyridinium for the transportation of sputum in tuberculosis bacteriology. Tubercle 1979;60:31-6. 25. Yeager H Jr, Lacy J, Smith LR, LeMaistre CA. Quantitative studies of mycobacterial populations in sputum and saliva. Am Rev Respir Dis 1967;95:998-1004. 70 Identification Test Techniques Identification Test Techniques Introduction Once an acid-fast isolate has been relegated to a subgroup on the basis of pigment production and growth rate, specific identification is accomplished by the performance of a battery of in vitro tests. Final identification should never be based upon a single test or characteris- tic because individual strains may deviate from the expected results of a given species. The reaction pattern of the isolate should be com- pared to the hypothetical median reaction patterns of known species. If an unknown matches any one pattern exactly, the identification may be considered definitive for that species. Such absolute correla- tion of reaction patterns occurs in 85% to 95% of the cases, depend- ing on the laboratory. Should the reactions of the unknown not match any indicated patterns, repeat the tests at least once; a second failure would suggest that the organism must receive more detailed study, generally in a larger reference laboratory. Many of the isolation and biochemical test media described in this manual are now available from a number of sources. The list below changes frequently, but it may be used as a guide; check with manu- facturers and journal advertisements for updates. As with any com- mercial product, it is best to compare these against the isolation media or test method you used before making the change to commer- cial suppliers. The availability of such commercial products, provided they are subject to good quality control, should ultimately lead to better standardization of methods in the mycobacteriology laboratory. The following prepared media are currently available from the indi- cated companies: Commercial Companies (Key to identification follows) Plated Media Dubos Oleic Acid Albumin Agar 3,5 MacConkey w/o Crystal Violet 5 Middlebrook 7H-10 Agar 1,3,4,5 Middlebrook 7H-11 Agar 2,345 Mitchison 7H-11 Selective Agar 5 71 Commercial Companies (Key to identification follows) Tubed Media American Trudeau Society - slant 1,2,3,4,5,6 American Trudeau Society w/5% NaCl - slant 2 Arylsulfatase Broth - 3-day test 5,6 Arylsulfatase Broth - 2-week test 5,6 Arylsulfatase Agar - butt 5,6 Dubos Oleic Acid Agar - butt 35 Dubos Tween Albumin Broth 3,5,6 Lowenstein-Jensen - slant 1,2,3,4,5,6 Lowenstein-Jensen - butt 1,2,3,4,5,6 Lowenstein-Jensen w/5% NaCl - slant 1,2,3,4,5 Lowenstein-Jensen Gruft (w/Penicillin, Nalidixic Acid) 2,3,4,5,6 Lowenstein-Jensen - Mycobactocel (w/Cycloheximide, Lincomycin, Nalidixic Acid) - slant 1,4,5 Lowenstein-Jensen (W/RNA) - slant 2 Lowenstein-Jensen (W/RNA) - butt 2 Middlebrook 7H-9 Broth w/Tween 80 1,2,4,5,6 Middlebrook 7H-10 Agar - slant 1,3,4,5,6 Middlebrook 7H-11 - slant 2,3,4,5,6, Mitchision 7H-11 Selective Agar - slant 5 Nitrate Substrate Broth 5 Pyrazinamidase Agar - butt 5 TCH (10 mcg/ml) Agar - slant 5 Tellurite Reduction (same as 7H-9 Broth w/Tween 80) 5 Urea Broth 5 Wayne Sulfatase Agar 1.4 Reagents Ferric Ammonium Citrate (20%) 5 NAC (mucolytic agent) N-acetyl-L- cysteine 5 OADC (Oleic Acid Albumin Dextrose Catalase) Enrichment 1,3,4,5 Potassium Tellurite (0.2%) 5 Sputagest 50 (mucolytic agent) Cleland’s Reagent-Dithiothreitol 5 TB Base Digestant (Sodium Hydroxide/ Sodium Citrate) 5 Tween 80 (10%) Catalase Test 5 Tween 80 Hydrolysis Substrate Concentrate 3,6 72 Stains Kinyoun Carbolfuchsin Ziehl-Neelsen Carbolfuchsin Decolorizer (3% Acid Alcohol) Decolorizer (Truant-Moore) Brilliant Green Counterstain Malachite Green Counterstain Methylene Blue Counterstain Auramine-0 Fluorescent Auramine-Rhodamine Fluorescent Potassium Permanganate Fluorescent Counterstain Commercial Companies (Key to identification follows) 2,345 2,345 23,45 3,5 3,5 3,5 2,345 34,5 3,5 3,5 KEY 1. Bioquest 4. P.O. Box 243 Cockeysville, MD 21030 2. Carr-Scarborough 5. Microbiologicals, Inc. P.O. Box 1328 Stone Mountain, GA 30086 3. Difco Laboratories 6. P.O. Box 1058A Detroit, MI 48232 Gibco Laboratories 3175 Staley Road Grand Island, NY 14072 Remel, Regional Media P.O. Box 14428 12076 Sante Fe Drive Lenexa, KS 66215 Scott Laboratories 771 Main Street Fiskeville, RI 02823 A. ARYLSULFATASE (4, 5) Principle Arylsulfatase is an enzyme capable of hydrolyzing the bond between the sulfate group and the aromatic ring structure in compounds with the general formula R-OSO3H, where R represents an aromatic ring compound. When the substrate potassium phenolphthalein disulfaté is incorporated into the growth medium, the presence of free phenol- phthalein liberated by enzymatic hydrolysis is indicated by a red color that develops when alkali is added. 73 Arylsulfatase activity is limited almost solely to the genus Myco- bacterium. Since most mycobacteria possess this enzyme, the test conditions must be varied to help identify the various mycobacteria. The 3-day test facilitates identification of the potentially pathogenic M. fortuitum complex, M. xenopi, and M. triviale, whereas the 14-day test aids in the identification of certain slowly growing mycobacteria, such as M. marinumand M. szulgai. Methods 1. Centers for Disease Control (CDC) Method (1, 2) The CDC method utilizes a liquid substrate medium for both the 3-day and the 14-day tests. eo Media and Supplies Phenolphthalein disulfate, tripotassium salt Dubos Tween-albumin or Middlebrook 7H-9 broth Middlebrook Albumin Dextrose Catalase (ADC) enrichment Sodium carbonate, Na,CO3 2N Sterile distilled water e Preparations Enzyme Substrate Stock Solution Dissolve 2.6 g of phenolphthalein disulfate, tripotassium salt, in 50 ml of distilled water to make a 0.08 M solution. Sterilize by filtration through a 0.22 wm pore size membrane filter and store at 5°C. Liquid medium Use either Dubos Tween-albumin or Middlebrook 7H-9 liquid medium, both of which are commercially available and should be prepared according to label directions. Prepare two flasks, each with 180 ml of the basal medium, and sterilize by autoclaving for 15 min- utes at 121°C. After cooling to room temperature, aseptically add 20 ml of commercial Middlebrook ADC enrichment to each flask to pro- vide a total volume of 200 ml. For the 3-day test medium, asceptically add 2.5 ml of the 0.08 M enzyme substrate stock to one flask of 200 ml of complete liquid medium to provide a 0.001 M substrate. For the 14-day test medium, aseptically add 7.5 ml of the 0.08 M enzyme substrate stock to the other flask of 200 ml of complete liquid medium to provide a 0.003 M substrate. Aseptically dispense each working solution in 2-ml amounts into sterile 16 x 125-mm screwcap test tubes and store at room tempera- ture or in the refrigerator. To prevent confusing the 3-day with the 14-day test medium, use color coded tubes. 74 2N Sodium carbonate Dissolve 10.6 g of anhydrous, Na,CO3; in 100 ml of distilled water. Sterilize by membrane filtration, 0.22 um pore size, if long-term stor- age (and contamination) is a problem. e Controls A tube of uninoculated substrate medium is used as a negative control for color change. A tube of substrate medium inoculated with M. fortuitum is used as a positive control. A tube of substrate medium inoculated with M. smegmatis or M. avium is used as a negative control for the 3-day test. Strains of M. tuberculosis or M. avium known to be negative in the 14 day test may be used to control the 14-day arylsulfatase test. e Procedure (1) Label tubes of 3-day and 14-day test substrates with speci- men numbers. (2) Inoculate both tests with 0.1 ml of a 7-day liquid culture of the unknown organism or a spadeful of organisms taken from an actively growing subculture on solid medium. (3) Incubate at 35 to 37°C. (4) After 3 days of incubation, add no more than 6 drops of 2N sodium carbonate to the 0.001 M substrate tube. (5) After 14 days of incubation, add no more than 6 drops of 2N sodium carbonate to the 0.003 M substrate tube. e Results and Interpretation A positive reaction = immediate color change to pink or red after addition of carbonate solution (see color plate 4.) A negative reaction = no color change. Positive results may be compared with a set of color standards (color plate 5) and the intensity of color recorded from + to 5+. e Color Standards for CDC Arylsulfatase Test Media and Supplies 0.067 M disodium phosphate, Na,HPO, Phenol red, 0.1% Screwcap test tubes, 16 x 125-mm 75 fel= b, X COLOR PLATE 4. Arylsulfatase—CDC method. | ] | 3 3+ 4+ 5+ COLOR PLATE 5. Color standards for arysulfatase test. Preparations 0.067 M DISODIUM PHOSPHATE Dissolve 9.47 g of anhydrous Na,HPO, in 1000 ml of distilled water. 0.1% PHENOL RED Dissolve 0.1 g of phenolsulfonphthalein (sodium salt) in 100 ml of distilled water. Autoclave and store in the refrigerator. Prepare six screwcap test tubes (16 x 125 mm) as follows: 0.067 M Tube Label Tube No. Na,HPO, 0.1% Phenol Red & Reading 1 4 ml 10 drops 0.1% stock 5+ 2 4 ml 1 ml, from tube 1, mix 4+ 3 3 ml 2 ml, from tube 2, mix 3+ 4 3 ml 2 ml, from tube 3, mix 2+ 5 3 ml 2 ml, from tube 4, mix 1+ 6 3 ml 2 ml, from tube 5, mix de 76 Discard 2 ml from tube 6. Autoclave all tubes, seal, and store at 5°C. Color standards usually retain color for 6 months. Precautions Always check a few tubes of the prepared, uninoculated substrate medium for free phenolphthalein by adding a few drops of 2N sodium carbonate. The substrate should remain colorless. The formation of a pink color indicates the presence of free phenolphthalein. This could give false-positive results. If this occurs, it indicates premature break- down of either the test medium or the powdered phenolphthalein disulfate salt. Discard the bad batch of medium and test the pure pow- der for free phenolphthalein by dissolving a small quantity of the powder in a mimimal amount of water. Add a few drops of 2N sodium carbonate. The formation of a pink color indicates free phenolphtha- lein, which will give positive reactions under actual test conditions. Sufficient purification may be obtained by dissolving 3 to 5 g of phe- nolphthalein disulfate in a minimal amount of distilled water and reprecipitating the salt by adding an excess of ethanol. Filter off the precipitated disulfate salt on a Buchner funnel. Wash the precipitate several times in fresh ethanol and allow to dry before bottling. 2. Wayne’s Phenolphthalein Sulfatase Test (3) The Wayne modification utilizes an unenriched agar base medium that provides consistent results only for the 3-day test. eo Media and Supplies Dubos oleic agar base Glycerol Phenolphthalein disulfate, tripotassium salt Sodium carbonate, Na,COs3, 2N Flat-bottomed screwcap vials, 18 x 160-mm e Preparations Solid Medium Add 1 ml of glycerol and 65 mg of phenolphthalein disulfate, tripotassium salt to 100 ml of melted Dubos oleic agar base. Dispense 2-ml amounts into 18 x 160-mm flat-bottomed screwcap vials (or 1-0z. screwcap prescription bottles) and autoclave. Place tubes in an upright position until agar hardens. Wayne sulfatase agar containing the substrate salt is commercially available in powder form. 2N Sodium carbonate Dissolve 10.6 g of anhydrous Na,CO3; in 100 ml of distilled water. 77 e Controls An uninoculated tube of substrate medium serves as a negative medium control. A tube of substrate medium inoculated with M. fortuitum serves as a positive control. A tube of substrate medium inoculated with M. smegmatis or M. avium serves as a negative control. e Procedure (1) Prepare a barely turbid suspension of the test organism in sterile water. (2) Inoculate butt surface of substrate agar with 1 drop of this suspension. (3) Incubate at 35 to 37°C. (4) After 3 days of incubation, add 1 ml of sodium carbonate solution and observe for color change. e Results and Interpretation A positive reaction = formation of a pink-to-red band on the surface of the agar within 30 minutes after adding the carbonate indicates release of free phenolphthalein. A negative reaction = no color change after 30 minutes (color plate 6). REFERENCES 1 2. ly RA ~ Cc Cc * COLOR PLATE 6. Wayne's phenolphthalein sulfatase test. Kubica GP, Ridgon AL. The arylsulfatase activity of acid-fast bacilli. Ill. Preliminary investigation of rapidly growing acid-fast bacilli. Am Rev Respir Dis 1961;83:737-40. Kubica GP, Vestal AL. The arylsulfatase activity of acid-fast bacilli. |. Investigation of stock cultures of acid-fast bacilli. Am Rev Respir Dis 1961;83: 728-32. . Wayne LG. Recognition of Mycobacterium fortuitum by means of the 3-day phenol- phthalein sulfatase test. Am J Clin Pathol 1961;36:185-97. . Whitehead JEM, Morrison HR, Young L. Bacterial arylsulfatase, Biochem J 1952;51: 585. . Whitehead JEM, Wildy P, Engbaek HC. Arylsulfatase activity of mycobacteria. J Pathol Bacteriol. 1953;65:451-60. 78 B. Carbon Sources (1, 2) Principles The ability to grow on medium containing either sodium citrate, mannitol, or inositol as a sole source of carbon in the presence of ammoniacal nitrogen proves helpful in identifying some rapidly grow- ing mycobacteria. Method eo Media and Supplies Ammonium sulfate, (NH), SO4 Monopotassium phosphate, KH,PO,4 Magnesium sulfate (heptahydrate), MgSO,4-7H,0 Agar Distilled water Potassium hydroxide (KOH), 10% Hydrochloric acid (HCI), 10% Sodium Citrate Mannitol Inositol Screwcap tubes, 20 x 150-mm e Preparations Basal Medium Dissolve the following ingredients in 950 ml of distilled water: 2.4 g (NH4)>S04, 0.5 g KH,PO4, 0.5 g MgS0,4.7H,0. Dissolve ingredients with gentle heating and stirring. Adjust pH (as indicated below) with either 10% KOH or 10% HCI, and add 20 g of purified agar. Boil to dissolve and sterilize by autoclaving at 121°C for 20 minutes. For Citrate Test Medium Adjust the pH of 950 ml of basal medium to 7.0; autoclave-sterilize and cool the medium to 56°C in a water bath. Dissolve 5.6 g of sodium citrate in 50 ml of distilled water, sterilize by membrane filtration, and add aseptically to sterile, tempered basal medium. For Mannitol Test Medium Adjust the pH of 950 ml of basal medium to 7.2; autoclave-sterilize and cool the medium to 56°C in a water bath. Dissolve 5.0 g of manni- tol in 50 ml of distilled water, sterilize by membrane filtration, and add aseptically to sterile, tempered basal medium. For Inositol Test Medium Adjust the pH of 950 ml of basal medium to 7.2; autoclave-sterilize and cool the medium to 56°C in a water bath. Dissolve 5.0 g of inositol 79 in 50 ml of distilled water, sterilize by membrane filtration, and add aseptically to sterile, tempered basal medium. For Control Medium Adjust the pH of the basal medium to 7.2 before autoclaving. Use 1000 ml of distilled water, since no carbon source is added. Tube all of the above media in 8-ml amounts in sterile, 20 x 150-mm screwcap tubes. Allow media to solidify at a slant. e Controls The test provides its own control. e Procedure (1) Using a 7-day old 7H-9 broth culture of the test organism, make serial tenfold dilutions in sterile saline until no turbid- ity is visible. (2) Use this last suspension to inoculate 0.1 ml onto each of the carbon source media and one control slant. (3) Incubate all slants at 28°C for 2 weeks. e Results and Interpretation Positive = growth on the test medium (citrate, inositol, or mannitol) but no growth on the control medium. Negative = no growth on control or test media. REFERENCES 1. Silcox VA, Good RC, Floyd MM. Identification of clinically significant Mycobacterium fortuitum complex isolates. J Clin Microbiol 1981; 14:686-91. 2. Tsukamura M. Identification of mycobacteria. Obu, Aichi-Ken, Japan: Research Laboratory, National Sanatorium Chubu Chest Hospital, 1975;1-75. C. CATALASE Principle Catalase is an intracellular, soluble enzyme capable of splitting hydrogen peroxide into water and oxygen, 2H,0, = 2H,0 + O,. The oxygen bubbles into the reaction mixture to indicate catalase activity. Virtually all mycobacteria possess catalase enzymes, except for cer- tain isoniazid-resistant mutants of M. tuberculosis and M. bovis. Stud- ies have shown that mycobacteria possess several kinds of catalase that vary in heat stability. The quantitative differences in catalase activity demonstrated by intact cells in the semiquantitative test, and the differences in heat stability detected by the 68°C catalase test have proved to be useful taxonomic tools. All mycobacteria fall into 80 one of the following groups with respect to their catalase activity: (a) those devoid of catalase; (b) those producing less than 45 mm of bubbles in the semiquantitative test; (c) those producing more than 45 mm of bubbles in the semiquantitative test; and (d) those losing catalase activity when heated to 68°C for 20 minutes. Methods 1. Semiquantitative Catalase Test (1, 3) This test divides the mycobacteria into two groups, those pro- ducing less than 45 mm of bubbles and those producing more than 45 mm of bubbles. Generally, M. kansasii, M. simiae, most scotochromogens, the nonphotochromogenic saprophytes, and the rapid growers produce more than 45 mm of bubbles in this test. Among those that produce less than 45 mm of bubbles are M. tuberculosis, M. marinum, M. avium complex, M. xenopi, and M. gastri. eo Media and Supplies Lowenstein-Jensen or other egg-base medium, prepared as butts or deeps Tween 80, 10% Hydrogen peroxide (H,0,), 30% Screwcap test tubes, 20 x 150-mm Preparations Lowenstein-Jensen butt (or deep) Dispense Lowenstein-Jensen (L-J) or other egg-base medium in 5-m| amounts into 20 x 150-mm screwcap test tubes and inspissate at 85°C for 50 minutes with tubes in an upright position to provide a butt or deep of solid medium. L-J butts also are commercially available. 10% Tween 80 Mix 10 ml of Tween 80 with 90 ml of distilled water and autoclave for 10 minutes at 121°C. Swirl solution immedi- ately after autoclaving and during cooling to resolubilize Tween. Store at 5°C. 30% Hydrogen peroxide (Superoxol, Merck), store at 5°C Just before use, mix equal parts of 10% Tween 80 and 30% hydrogen peroxide. Prepare approximately 1.0 ml of mixture for each test. Caution: Wear rubber or plastic gloves and protective eye shield when handling Superoxol. 81 Controls An uninoculated tube of medium is used as a negative control. For the low catalase control, inoculate either M. gastri or M. avium complex onto a tube of butt medium. Mycobacterium gordonae or M. terrae complex may be used for the high catalase control. Procedure (1) (2) (3) (4) Inoculate the butt medium surface with 0.1 ml of a 7-day-old liquid culture of the test organism or a loopful of growth from an actively growing slant. Incubate tubes with caps loose for 2 weeks at 35 to 37°C. After incubation, add 1.0 ml of freshly prepared Tween- peroxide mixture, replace caps loosely, and allow to stand at room temperature for 5 minutes before measuring. Measure in millimeters the height of the column of bubbles above the medium surface. Results and Interpretation Divide the test organisms into two groups: those that produce more than 45 mm of bubbles and those that pro- duce less than 45 mm of bubbles (color plate 7). COLOR PLATE 7. Semiquantitative catalase test. Heat Stable Catalase Test (at pH 7/68°C) (2) Some mycobacteria lose catalase activity when suspended in pH 7 buffer and heated to 68°C for 20 minutes. Included in this group are M. tuberculosis, M. bovis, M. gastri, and M. haemo- philum. This “hot catalase’ test is especially valuable in identify- ing strains of M. tuberculosis that give weakly positive or nega- tive niacin tests. All slow-growing nonchromogenic mycobacteria should be subjected to this test. 82 eo Media and Supplies 0.067 M Phosphate buffer, pH 7 Tween 80, 10% Hydrogen peroxide, 30% Screwcap tubes, 16 x 125-mm e Preparations 0.067 M Phosphate buffer, pH 7 Buffer solutions are prepared by mixing two stock phos- phate solutions to give the desired pH. Stock solutions: (1) 0.067 M disodium phosphate Dissolve 9.47 g of anhydrous Na,HPO, in water to make one liter (1000 ml). (2) 0.067 M monopotassium phosphate Dissolve 9.07 g of KH,PO,4 in distilled water to make one liter (1000 ml). To prepare a pH 7 buffer solution, mix 61.1 ml of #(1) stock solution with 38.9 ml of #(2) stock solution. Check on pH meter. 10% Tween 80 and 30% hydrogen peroxide are prepared as described in the “Semiquantitative Method." e Controls Always include a positive and negative control. Use MW. tuberculosis or M. gastri for a negative control, and M. gordonae or M. terrae complex as a positive control. e Procedure (1) Suspend several spadesful of growth from a culture slant of the test organism in 0.5 ml of 0.067 M phos- phate buffer, pH 7, contained in a 16 x 125-mm screw- cap tube. (2) Incubate this suspension in a 68°C water bath (or tem- perature block heater) for 20 minutes. Time and tem- perature are critical. (3) Cool the suspension to room temperature before add- ing 0.5 ml of the Tween-peroxide mixture. Recap tubes loosely. (4) Observe for the formation of bubbles. Hold negative tubes for 20 minutes before discarding. e Results and Interpretation A positive test is indicated by the formation of bubbles. No bubbles indicate a negative test (color plate 8). On rare occasions, bubbles may be seen rising from the sedimented cells in such small quantity that foam does not form at the 83 surface of the fluid. This is still recorded as a positive reaction. e Precautions Do not shake the tubes! Tween 80 alone may form bub- bles when shaken, resulting in false-positive reaction. COLOR PLATE 8. Heat stable catalase test. REFERENCES 1. Kubica GP, Jones WD Jr, Abbott VD, Beam RE, Kilburn JO, Cater JC Jr. Differential identification of mycobacteria. |. Tests on catalase activity. Am Rev Respir Dis 1966;94:400-5. 2. Kubica GP, Pool GL. Studies on the catalase activity of acid-fast bacilli. I. An attempt to subgroup these organisms on the basis of their catalase activity at different temperatures and pH. Am Rev Respir Dis 1960.81:387-91. 3. Wayne LG, Doubek JR. Diagnostic key to mycobacteria encountered in clinical laboratories. Appl Microbiol 1968;16:925-31. D. Growth Rate (1,2) Principles Mycobacteria are commonly grouped according to their rate of growth. The slow growers take more than 7 days to appear on culture media, whereas rapid growers are fully mature in less than 7 days. As a result of the harsh digestion-decontamination procedures used for primary isolation, some rapid growers may take as much as 3 weeks to appear on the primary culture medium. Consequently, growth rate studies should always be determined by using subcultures diluted sufficiently to yield isolated colonies on whatever solid medium (usually egg-base) is inoculated. 84 Method eo Media and Supplies Middlebrook 7H-9 broth Screwcap test tubes, 20 x 150-mm Sterile 9-m| water blanks for dilutions Egg- or agar-base medium in tubes or plates. e Preparations Middlebrook 7H-9 broth from commercial base Suspend 4.7 g of dehydrated medium in 900 ml of distilled water containing 2 ml of glycerol or 0.5 ml of Tween 80. Do not use glycerol and Tween 80 together. Autoclave at 121°C for 15 minutes. Cool to 45°C before aseptically adding 100 ml of ADC enrichment. Aseptically dispense 5-m| amounts into sterile 20 x 150-mm screwcap test tubes. eo Procedure (1) (2) (3) (4) (5) Determine purity of the primary isolation culture by smear examination. Aseptically pick several colonies and inocu- late into 7H-9 broth. Incubate culture at 35°C for 5 to 7 days with daily shaking to enhance growth. Check broth culture for purity, using smear examination. After incubation, dilute the bacterial suspension sufficiently (“high dilution”) to yield isolated colonies. This may require several tenfold dilutions. Selection of the “’high dilution” is dependent upon the turbidity of the broth culture (see chart below). Broth Dilutions to be made Consistency Low Medium High Very turdid 104 10-5 10° Moderately turbid 10°20r 103 102%or 10° 10%or 10% Slightly turbid undil. or 10°" 107" or 102 1020r 10°73 Until experience is gained in choosing the proper dilu- tion to obtain isolated colonies, tubes of media should also be inoculated with the next lower or “medium dilution.” The chart lists the recommended dilutions, dependent on broth consistency. Inoculate 0.1 ml of each of these tenfold dilutions onto either egg- or agar-base medium. Incubate at 35°C or at the optimum temperature for those organisms with a more restricted temperature growth range. 85 (6) Examine inoculated media after 5 to 7 days and weekly thereafter for grossly visible colonies. e Results and Interpretation Record the growth rate as the number of days required for the appearance of mature, grossly visible colonies. Rapid grower - less than 7 days. Slow grower - more than 7 days. REFERENCES 1. Kubica GP, Gross WM, Hawkins JE, Sommers HM, Vestal AL, Wayne LG. Laboratory services for mycobacterial disease. Am Rev Respir Dis 1975;112:773-87. 2 Runyon EH, Karlson AG, Kubica GP, Wayne LG. Mycobacterium (revised by Som- mers HH, McClatchy JK). In: Lennette EH, Balows A, Hausler WJ Truant JP, eds. Manual of clinical microbiology. 3rd ed. Washington, DC.: American Society for Microbiology, 1980. E. Iron Uptake Principle The iron uptake test is used to detect those mycobacteria capable of converting ferric ammonium citrate to an iron oxide. This “rusting” colors both the colonies and the medium a reddish brown. Iron uptake is useful in distinguishing M. chelonae, commonly negative, from M. fortuitum and most other rapid growers that are positive. Slow growers and most M. flavescens are not capable of accumulating iron oxides. Methods 1. Modified Method of Tison et al. (2) Add a final concentration of 2.5% ferric ammonium citrate into Lowenstein-Jensen egg medium before inspissation. Inspissate and check for sterility as usual. The final medium appears “dirty green.” eo Media and Supplies Lowenstein-Jensen egg medium Salt solution Potato flour Malachite green, 2% Whole eggs Ferric ammonium citrate 86 e Preparation Whole Eggs Scrub fresh eggs, not more than 1 week old, with a hand brush and a soapy solution. Soak the eggs for 30 minutes in the soapy solution. Rinse thoroughly with running water and soak the eggs for 15 minutes in 70% alcohol. Scrub hands well with soap and finally rinse in 70% alcohol before breaking the eggs into a sterile flask. Homogenize the eggs by hand shaking, then filter through four layers of sterile gauze into a sterile graduated cylinder. Salt Solution Combine: Monopotassium phosphate (anhydrous), Magnesium sulfate-7 H,0, MgS0,-7H,0 0.24 g Magnesuim citrate 06g Asparagine 36g Glycerol (reagent grade) 12.0 ml Distilled water 600.0 ml Malachite Green Dissolve 0.4 g of malachite green in 20 ml of distilled water to obtain a 2% aqueous solution. Final Medium « Add 30 g of potato flour to the above salt solution. e Autoclave at 121°C for 30 minutes. « Cool to room temperature before adding 20 ml of the freshly prepared 2% malachite green and 1000 ml of homogenized whole eggs. « Add a final concentration of 2.5% (wt/vol.) ferric ammo- nium citrate. « Mix and pour into a sterile aspirator bottle or funnel with a bell attachment (test tube filling device) and dispense. « Place approximately 6 to 8 ml of medium into each 20 x 150-mm sterile screwcap test tube. « Slant tubes and coagulate by inspissation at 85°C for 50 minutes. « Incubate at both 25 and 37°C for 48 hours each as a sterility check. « Medium may be stored in the refrigerator for several months if caps are tightly closed to prevent evapora- tion. e A batch of L-J medium without iron citrate (control) should be made at the same time. 87 eo Controls Perform the test by inoculating both an iron citrate- containing slant and an iron citrate-free slant of Lowenstein- Jensen (L-J) medium. The iron citrate-free slant is a control to verify that the organism will grow on L-J medium. Use M. fortuitum as a positive control and M. chelonae as a negative control. Compare the pigmentation of the colonies observed on L-J medium without ferric ammo- nium citrate to that of the same culture on L-J with iron citrate added. e Procedure (1) Inoculate the media with one drop of a barely turbid suspension of the organism being tested. (2) Incubate cultures at 28°C for 2 weeks with caps loose. (3) Examine weekly for 3 to 4 weeks before recording as negative. e Results and Interpretation A positive reaction = colonies turn rusty brown because of the uptake of iron. The medium may also be tan to rusty brown in color (color plate 9). A negative reaction = no growth on iron citrate medium but growth on control L-J medium. OR A negative reaction = growth on both media without visi- ble rusting of the colonies. On occasion, the colonies become tan, and a beige color may develop around the edge of the slant; this reaction is recorded as =. Some strains of M. chelonae (designated “turtle or turtle-like’’) exhibit this intermediate reaction. + a—— ATTN . ¥ % oO he: ” COLOR PLATE 9. Iron uptake. 2. Modified Method of Szabo and Vandra (Wayne) (1, 3) This method requires that ferric ammonium citrate be added to the L-J slant after visible growth is observed. eo Media and Supplies Lowenstein-Jensen egg medium Ferric ammonium citrate e Preparations Lowenstein-Jensen egg medium Lowenstein-Jensen slants are commercially available or may be prepared as described in Method 1, except that the ferric ammonium citrate must not be incorporated. Ferric ammonium citrate Dispense 20% aqueous solution of ferric ammonium citrate in 5 to 8-ml amounts and autoclave. Discard if the solution precipitates or becomes cloudy and prepare fresh. eo Controls Inoculate one L-J slant with M. fortuitum (positive control) and another with M. chelonae (negative control). e Procedure (1) Inoculate two L-J slants with one drop of a barely turbid suspension of the test organism. (2) Incubate in slanted postion with caps loosened until colonies are grossly visible. (3) Add sterile ferric ammonium citrate to only one of the L-J tubes. The other is used as a negative color control. Add one drop citrate for each milliliter Lowenstein- Jensen medium (usually 6 to 8 drops per slant). (4) Reincubate cultures at 28°C and examine weekly for 3 weeks. e Results and Interpretation A positive reaction = colonies appear rusty brown because of the iron being taken up. A negative reaction = pigmentation of colonies remained the same as that observed on L-J medium that did not receive citrate solution. REFERENCES 1. Szabo |, Vandra E. Mycobacterium minetti (Penso et al., 1952) bacteriological and epidemiological observations. Acta Microbiol Acad Sci Hung 1963;10:215-23. 2. Tison F, Tacquet A, Devulder B. Un test simple d’étude mycobactéries: la transfor- mation du citrate de fer ammoniacal. Ann Inst Pasteur 1964;106:797-801. 3. Wayne LG, Doubek JR. Diagnostic key to mycobacteria encountered in clinical laboratories. Appl Microbiol 1968;16:925-31. 89 F. Growth on MacConkey Agar Without Crystal Violet (3) Principle The MacConkey agar test is of value in the taxonomic separation of the rapidly growing acid-fast bacilli (1). The potentially pathogenic rapid growers of the M. fortuitum complex usually grow on Mac- Conkey agar and sometimes reveal a color change in the medium within 5 to 11 days, whereas the commonly saprophytic species are inhibited by this medium. It should be noted that about 25% of the M. smegmatis strains have been found to grow on MacConkey agar without crystal violet (2). To obviate any diagnostic difficulties posed by the rare isolation of M. smegmatis from humans, the 3-day arylsulfatase test may be used to separate M. smegmatis from the M. fortuitum complex. Mycobacterium fortuitum and M. chelonae are positive, whereas M. smegmatis is rarely positive in the 3-day arylsulfatase test (4). Method eo Media and supplies MacConkey agar without crystal violet Spray-Fisher petri dish turntable Plastic petri dishes, 15 x 100-mm Loop, 3-mm e Preparations MacConkey agar without crystal violet Prepare commercially available MacConkey agar without crys- tal violet according to directions on the label. Pour 20 ml of medium into each 15 x 100-mm petri dish. Culture Cultures to be tested should be grown for 7 days at 28°C in enriched Middlebrook 7H-9 liquid medium. e Controls Always include an uninoculated control plate along with a known negative organism, e.g., M. phlei, and a known positive organism, e.g., M. fortuitum. e Procedure (1) Place a MacConkey agar plate on the turntable, remove plate cover, spin the turntable, touch a 3-mm loopful of a 7-day broth culture to the center of the spinning plate, and gradually move the loop to the periphery of the plate. This produces a spiral of inoculum. By matching the movement of the loop with that of the spinning plate, it is possible to produce 6 to 12 spirals on the plate. An alternative method 90 is to streak plates as for isolated colonies, but results by this method are not as distinctive. (2) Incubate plates at 28°C in an open petri dish cannister. Do not incubate in a CO, incubator. (3) Examine plates at 5 and 11 days. e Results and Interpretation Most strains of M. fortuitum and M. chelonae grow along the full length of the spiral (color plate 10). A color change in the medium may or may not be evident but is not as important as the extent of growth. When a very heavy inoculum is used, some commonly negative saprophytic rapid growers may exhibit growth at the very beginning of the spiral. COLOR PLATE 10. Growth on MacConkey without crystal violet. REFERENCES 1. Jones WD Jr, Kubica GP. The differential typing of certain rapidly growing mycobacteria based on their sensitivity to various dyes. Am Rev Respir Dis 1963;88:355-9. 2. Jones WD Jr, Kubica GP. The use of MacConkey’s agar for the differential typing of Mycobacterium fortuitum. Am J Med Technol 1964;30:187-90. 3. Kubica GP, Vitvitsky J. Comparison of two commercial formulations of the Mac- Conkey agar test for mycobacteria. Appl Microbiol 1974;27:917-9. 4. Silcox VA, Good RC, Floyd MM. Identification of clinically significant Mycobacterium fortuitum complex isolates. J Clin Microbiol 1981;14:686-91. G. Niacin Production (6, 9) Principle Niacin plays a vital role in the oxidation-reduction reactions that occur during metabolic syntheses in all mycobacteria (1). It functions as a precursor in the biosynthesis of coenzymes. Although all mycobacteria produce nicotinic acid, comparative studies have shown that, because of a blocked metabolic pathway, M. tuberculosis 91 accumulates the largest amount, and detection of this accumulated niacin is useful for the definitive diagnosis of this species (1, 6). The niacin test should not be used alone to identify M. tuberculosis because several other species (M. simiae, M. chelonae chemovar niacinogenes, and some strains of BCG) consistently yield positive results (1). This fact alone emphasizes the importance of performing the supportive tests of nitrate reduction and 68°C catalase to confirm the identity of M. tuberculosis. Cultures grown on egg medium yield the most consistent results in the niacin test. When unusual circumstances necessitate the use of cultures grown on 7H-10 or 7H-11 agar, use media supplemented with 0.1% potassium aspartate (5) or by extracting the agar-grown cultures for 2 hours at 37°C (7, 8). Before the niacin test is performed, cultures should (a) be checked for purity by microscopy, (b) be 3 to 4 weeks old on egg medium, and (c) have sufficient growth of 50 or more colonies. If cultures are still niacin negative at 4 weeks and if they have been handled aseptically, they may be reincubated for retesting when 6 weeks old; otherwise, a fresh culture should be used. Because mycobacteria excrete niacin into the growth medium, cultures with confluent growth may give a false-negative niacin reaction because the extracting fluid cannot con- tact the culture medium. When this occurs, expose the underlying medium surface by either scraping away or puncturing through some of the culture growth. The niacin test may be done either with chemical reagents or with commercially available paper strips. Regardless of the method used, niacin is usually detected by its reaction with a cyanogen halide in the presence of a primary amine. Comparative studies have shown the chemical reagents and paper strips to give comparable results. Methods 1. Niacin Test with Chemical Reagents (6, 9) eo Media and Supplies Ethanol, 95% Aniline, 4% Cyanogen bromide, 10% Sterile distilled water or 0.85% saline Sterile screwcap test tubes, 16 x 125-mm e Preparations 4% Aniline Add 4.0 ml of colorless aniline to 96.0 ml of 95% ethanol. Store in a brown bottle in the refrigerator. Discard if solu- tion turns yellow. 92 10% Cyanogen bromide Dissolve 5 g of cyanogen bromide in 50 ml of distilled water. Store in the refrigerator in a tightly capped brown bottle. Warm to room temperature to dissolve any precipi- tate formed upon cooling. Prepare small amounts of this reagent because cyanogen bromide is volatile and loses strength on storage. Weak solutions give false-negative results. See PRECAUTIONS below. 0.85% saline Dissolve 0.85 g of sodium chloride in 100 ml of distilled water. Sterilize by autoclaving. Controls Control the reagents by testing the extract from an unin- oculated tube of medium. The result should be negative. Use an extract from a known culture of M. tuberculosis as a positive control and an extract from an M. avium complex as a negative control. Procedure (1) Add 1.0 ml of sterile, distilled water or saline to the egg-base medium culture slant to be tested. (2) Place the tube horizontally so the fluid covers the entire surface of the medium. (3) Allow at least 15 minutes for the extraction of niacin. The extraction time may be longer when the culture has few colonies or is known to be a weak niacin producer. (4) Remove 0.5 ml of the fluid extract to a clean 16 x 125-mm screwcap tube. (5) Add 0.5 ml of the 4% aniline solution and 0.5 ml of 10% cyanogen bromide. (6) Observe the solution for the immediate formation of a yellow color (color plate 11). Results and Interpretation Positive = yellow color. Negative = no color change. Precautions Some of the difficulties and problems encountered in performing the niacin test with chemical reagents have already been mentioned. The following precautions should also be observed: (a) Aniline may change color on expo- sure to air and light; redistill if necessary or prepare a fresh 93 AA A COLOR PLATE 11. Niacin chemical test. solution. (b) Cyanogen bromide is a severe lacrimator and may be quite toxic if inhaled; handle in a chemical fume hood when preparing the solution and in a biological safety cabinet when testing cultures. (Serious illnesses have resulted from cyanogen bromide poisonings in laboratories that were testing large numbers of cultures for niacin pro- duction in class Il laminar flow BSCs that were vented back into the laboratory.) This, alone, is strong justification for venting all safety cabinets to the outside. (c) In acid solution, cyanogen bromide hydrolyzes to hydrocyanic acid, which is extremely toxic. Handle cyanogen bromide only in a well- ventilated safety cabinet and discard all reaction tubes into a disinfectant solution made alkaline by the addition of sodium hydroxide. Niacin Test with Paper Strips (4, 10) Paper test strips for the detection of niacin are commercially available (Difco and Remel). These have been evaluated (2, 3) and found to be comparable to the chemical reagents for sensi- tivity in detecting niacin production by mycobacteria. A paper- strip method obviates the need to prepare and store the unsta- ble chemicals used to demonstrate the presence of niacin. When using paper strips, always follow the manufacturer's directions. e Media and Supplies Reagent-impregnated paper strips Sterile, sealable test tubes (screwcap, cork, or rubber stop- per), 12 x 75-mm (or 13 x 75-mm) e Controls Same as chemical procedure. 94 eo Procedure (1) Prepare niacin extraction as in the previous chemical reagent procedure, steps 1 through 3. (2) Remove 0.6 ml of the culture extract to the test tube. (3) Insert the strip with the identification end up (an arrow may indicate which end to insert first) and immedi- ately seal the tube. Do not let middle of strip get wet. (4) Leave at room temperature for 15 to 20 minutes; occa- sionally agitate the tube without tilting it. (5) Observe the color of the liquid in the bottom of the tube against a white background. Disregard any color formation on the strip itself; this may occur because of oxidation of chemicals, especially at the top of the strip (color plate 12). (6) Before discarding the reaction tubes, neutralize the strips with 10% sodium hydroxide, or discard them into alkaline disinfectant. e Results and Interpretation Positive = development of yellow color in extract. Negative = no color development in extract. e Precautions Always check the expiration date of commercial paper strips. To prevent false-negative results, promptly reseal tubes after inserting paper strip; if tubes are left unsealed, the gas evolved as chemicals mix on the strip may escape into the atmosphere. COLOR PLATE 12. Niacin paper strip test. 95 REFERENCES 1. David HL. Bacteriology of the mycobacterioses. Washington, D.C: U.S. Govern- ment Printing Office, 1976. 2. DiSalvo AF, Lindler GN. Evaluation of a paper strip test for detection for niacin production by mycobacteria. Am J Clin Pathol 1970; 53:871-3. 3. Gangadharam PRJ, Droubi AJ. A comparison of four different methods for testing the production of niacin by mycobacteria. Am Rev Respir Dis 1971;104:434-7. 4. Kilburn JO, Kubica GP. Reagent-impregnated paper strips for detection of niacin. Am J Clin Pathol 1968;50:530-2. 5. Kilburn JO, Stottmeier KD, Kubica GP. Aspartic acid as a precursor for niacin synthesis by tubercle bacilli grown on 7H-10 agar medium. Am J Clin Pathol 1968;50:582-6. 6. Konno K. New chemical method to differentiate human-type tubercle bacilli from other mycobacteria. Science 1956;124:985. 7. Neimeister RP. A preliminary report on the technique of extracting niacin from Mycobacterium tuberculosis cultured on 7H-10 agar. Am Rev Respir Dis 1969; 100: 401-2. 8. Neimeister RP. Technique for extracting niacin from Mycobacterium tuberculosis cultured on 7H-10 and 7H-11 agars. J Clin Microbiol 1982; 15:971-2. 9. Runyon EH, Selin MJ, Harris HW. Distinguishing mycobacteria by the niacin test. Am Rev Tuberc 1959; 79:663-5. 10. Young WD Jr, Maslansky A, Lefar MS, Kronish DP. Development of a paper strip test for detection of niacin produced by mycobacteria. Appl Microbiol 1970;10: 939-45. H. Nitrate Reduction Tests (7) Principle The ability of some mycobacteria to reduce nitrate has proved valu- able in differential identification of some mycobacteria that possess such similar characteristics as colonial morphology, pigmentation, and growth rate (6). Virtanen (7) observed that the mycobacteria dif- fered quantitatively in their ability to reduce nitrate and that this reac- tion was influenced by the age of the culture, temperature, enzyme inhibitors, and hydrogen ion concentration. Most mycobacterial cul- tures to be tested for nitrate reduction should be examined 4 weeks after inoculation onto the subculture medium. Rapid growers that exhibit good growth may be tested after 2 weeks. Species that reduce nitrate are M. tuberculosis, M. kansasii, M. szulgai, M. flavescens, the M. terrae complex, and most rapid growers except M. chelonae (6). The test can be performed by using chemical reagents (either liquid or dry crystalline) or reagent-impregnated paper strips, or by combin- ing the nitrate reduction with the niacin test. For best results in both the classical and the paper-strip test, use a heavy suspension of the organisms. The combined test often yields more intense reactions and more reliable results in the tests for both niacin production and nitrate reduction. 96 Methods 1. Classical Method with Liquid Reagents (6, 7) Media and Supplies Sodium nitrate, NaNO; Monopotassium phosphate, KH,PO,4 Disodium phosphate, Na,HPO,4-12H,0 Hydrochloric acid, HCI Sulfanilamide N-naphthylethylenediamine dihydrochloride Distilled water Zinc dust Screwcap test tubes, 16 x 125-mm Water bath or constant-temperature block heater, 37°C Preparations Substrate To prepare 0.01 M sodium nitrate in 0.022 M phosphate buffer, pH 7.0 dissolve in order the following chemicals in 100 ml of distilled water: 0.085 g NaNO, 0.117 g KH,POy,, 0.485 g Na,HPO,4-12H,0. Sterilize by autoclaving. Reagent #1 Carefully add 50.0 ml of concentrated HCI to 50.0 ml of distilled water. NEVER ADD WATER TO ACID. Reagent #2 Dissolve 0.2 g of sulfanilamide in 100.0 ml of distilled water. Reagent #3 Dissolve 0.1 g of N-naphthylethylenediamine dihydrochlo- ride in 100.0 ml of distilled water. Store the substrate and reagents in the dark at 5°C. Dis- card the reagents if the color changes or a precipitate forms and prepare afresh. Controls Positive = M. tuberculosis (3+ to 5+) Negative = M. bovis (BCG) or M. intracellulare Negative = reagents without organisms. Procedure (1) Add 0.2 ml of sterile distilled water to a 16 x 125-mm screwcap tube. 97 (2) Use a sterile spade or applicator stick to emulsify in the water 2 spadesful of growth from a 4-week-old culture on Lowenstein-Jensen or some other egg-base medium. (3) Add 2.0 ml of the NaNOj substrate to the tube. (4) Shake by hand and incubate upright for 2 hours in a 37°C water bath. (6) Remove from the water bath. (6) Add one drop of reagent #1. (7) Add two drops of reagent #2. (8) Add two drops of reagent #3. (9) Examine immediately for a pink-to-red color (color plate 13). Note: A color standard may be used to help read the test, see page 102. Results and Interpretation Positive = May range from pale pink (+) to deep red (5+) when compared with the color standards. Only 3+ to 5+ are considered “positive.” Negative = No color. If no color develops, the test is either negative or the reduction has proceeded beyond nitrite. Add a small amount of powdered zinc to all negative tests. (a) If nitrate is still present, it will be catalytically reduced by the zinc, and a red color will develop, indicating a true negative. (b) If no color develops when zinc dust is added, the original reaction was positive, but the nitrate was reduced beyond nitrite. Repeat the test in this case to con- firm the observation. 5+ COLOR PLATE 13. Nitrate reduction test with liquid reagents. 98 Method with Crystalline Reagent (8) This new dry crystalline reagent, developed by Lampe (4) and evaluated by Warren et a1.(8), has proved to be reliable and sensitive in the detection of nitrate reduction by mycobacteria. It is easy to prepare, has a shelf-life of at least 6 months, and has the added advantage that only one “reagent” is needed to detect nitrate rather than the three liquid reagents used in the conven- tional chemical test. eo Media and Supplies I-tartaric acid Sulfanilic acid N-(I-Naphthyl)-ethylenediamine dihydrochloride Sodium nitrate substrate Screwcap test tubes, 16 x 125-mm Water bath or constant-temperature block heater, 37°C e Preparation Sodium nitrate substrate Prepare as described in previous “Classical Method.” Crystalline reagent Mix well 1 part sulfanilic acid, 1 part N-(I-Napthyl) -ethylenediamine dihydrochloride, and 10 parts I-tartaric acid. (The latter is much more crystalline than the other two chemicals and may have to be ground in mortor and pestle to ensure good admixture of reagents). Store in a dark bottle at room temperature. The dry chemical mixture will be nonuniform in texture and appearance. e Controls Same as ‘‘Classical Method.” e Procedure (1) May be performed as in “’Classical Method" procedure, steps (1) through (5) or as in “Combined Niacin/Nitrate Test” procedure, steps (1) through (5). (2) Use the tip of a spatula to add a small amount of crystalline reagent to test solution (the quantity of reagent is not critical). (3) Examine immediately for a pink-to-red color and com- pare to color standard. e Results and Interpretation Same as in previous “Classical Method." 99 Nitrate Paper-Strip Method (2, 5) The paper-strip test method yields most consistent results with those mycobacteria that vigorously reduce nitrate; inconsistent results are seen with species that are slow or weak reducers of nitrate. Because M. tuberculosis is a strong nitrate reducer, the paper-strip test may provide reliable test results with this species. Media and Supplies Commercially available reagent-impregnated nitrite test strips (Difco or Remel) Screwcap test tubes, sterile, 13 x 100-mm Water bath or constant-temperature block heater, 37°C Sterile saline, 0.85% Controls Positive = a strong nitrate reducer (M. tuberculosis). Negative = M. bovis (BCG) or M. intracellulare. Negative = an uninoculated tube containing reagents with- out organisms. Procedure (1) Add 1.0 ml of sterile saline to a sterile, 13 x 100-mm screwcap test tube. (2) Use a sterile spade or applicator stick to emulsify in the saline two spadesful of growth from a 4-week-old culture on egg-base medium. (3) Use flame-sterilized forceps and carefully insert a nitrite test strip (arrow indicates which end to insert first); do not let the strip contact any fluid on the side of the tube. (4) Cap the tube tightly and incubate in a vertical position at 37°C for 2 hours. (5) After the first hour of incubation, shake the tube gently without tilting. (6) After 2 hours of incubation, tilt the tube six times to wet the entire strip. (7) Allow the tube to remain slanted for 10 minutes with the liquid covering the strip. Results and Interpretation Positive = top portion of the strip changes to light or dark blue. Negative = no color change. Precautions Because the strips are sensitive to sunlight, excess heat, and moisture, they should be stored between 2 to 8°C in 100 the original container, tightly capped. Discard strips if they become discolored, for this indicates deterioration of reagent. Do not rely on results of nitrite test strips if the M. tuber- culosis control culture gives weak or negative reactions. Combined Niacin-Nitrate Test (3) The combined niacin-nitrate test is a modification of an earlier method of Kubica (3). This method is performed at CDC because it provides more intensely colored positive reactions in both the niacin and nitrate tests (color plate 14). The nitrate reduction test may be more intensely colored in this combined procedure because of the electron donors present in egg medium (1). eo Media and Supplies Same as in previous “Classical Method.” e Preparations Same as in “Classical Method.” e Controls Same as in “Classical Method.” eo Procedures (1) Use a 4-week-old culture on Lowenstein-Jensen or some other egg-base medium. (2) Add 2.5 ml of the buffered nitrate substrate directly to the test culture slant. (3) Dislodge some growth into the substrate with the pipette tip. (4) Place the tube upright in a 37°C water bath for 2 hours. (5) After incubation, remove 0.6 ml of the substrate solu- tion from the slant to a clean test tube and proceed with any desired niacin test. (6) Then, to the remaining substrate solution in the cul- ture slant tube, add the reagents for the nitrate reduc- tion test (use either liquid reagents or dry, crystalline reagent). When using liquid reagents, add them di- rectly to the reaction fluid. Do not let reagents run down the egg slant because they may be absorbed by the medium, resulting in weak or false-negative re- actions. (7) Verify all negative tubes by adding a small amount of zinc dust. e Results and Interpretation Positive = formation of a pink-to-red color. Negative = no color change. If no color develops, the test is either negative or the reduction has proceeded beyond 101 nitrite. Add zinc dust and interpret results as in “Classical Method.” Results may be compared to a set of color stan- dards for interpretation (see below). COLOR PLATE 14. Combined niacin-nitrate test. e Nitrate Reduction Standards Stock Solutions (1) 0.067 M disodium phosphate (9.47 g of anhydrous Na,HPO, per liter) (2) 0.067 M monopotassium phosphate (9.07 g of KH,PO, per liter) (3) 0.067 M trisodium phosphate (25.47 g of NazPO,4-12H,0 per liter) (4) 1% phenolphthalein (1 g in 100 ml 95% ethyl alcohol) (5) 1% brom thymol blue (1 g in 100 ml 95% ethyl alcohol) (5A) 0.01% bromthymol blue: prepare by mixing 1.0 ml of No. 5 above in 100 ml of distilled water. Working Buffer Solution Mix: Stock No. (1) 35 ml Stock No. (2) 5 ml Stock No. (3) 100 ml Preparation of Standards (1) Place eight clean test tubes (numbered 1-8) in a rack. Use the same size tubes as used to perform the nitrate reduction test. (2) Put2 ml ofworking buffer solution into tubes 2 through 8. (3) To 10 ml of working buffer solution, add 0.1 ml of stock No. (4) and 0.2 ml of stock No. (5A). (4) Add 2 ml of solution from step 3 to the tube num- bered 1. This is the 5+ color standard. 102 (5) To the tube numbered 2 in the series, add 2 ml of the solution from step 3. Mix well and transfer 2 ml to the next tube (#3). Continue to make serial dilutions of 2 ml, discarding 2 ml from the 8th tube. (6) The color standards: tube 1 = 5+ tube 2 = 4+ tube 3 = 3+ tube 5 = 2+ tube 6 = 1+ tube 8 = + These colors should range from pink (+) to purplish red (5+) (color plate 15). Discard tubes 4 and 7. (7) Autoclave tubes, seal, and store at 5°C. +m 2+ 3 ar ox COLOR PLATE 15. Nitrate color standards. REFERENCES 1. Bonicke R, Juhasz SE, Piemer U. Studies on the nitrate reductase activity of mycobacteria in the presence of fatty acids and related compounds. Am Rev Respir Dis 1970;102:507-15. 2. Difco Laboratories. Detroit: Bulletin No. 3183. 3. Kubica GP. A combined niacin-nitrate reduction test for use in the identification of mycobacteria. Acta Tuberc Pnemol Scand 1964; 45:161-7. 4. Lampe AS. Nonliquid reagent for detecting nitrate reduction. J Clin Microbiol 1981;14:452-4. 5. Quigley HJ, Elston HR. Nitrate test strips for detection of nitrate reduction by mycobacteria. Am J Clin Pathol 1970;53:663-5. 6. Runyon EH, Karlson AG, Kubica GP, et al. Mycobacterium. In: Lennette EH, Spaulding EH, Truant JP, eds. Manual of clinical microbiology, 2nd ed. Washington, D.C.: American Society for Microbiology, 1974. 7. Virtanen S. A study of nitrate reduction by mycobacteria. Acta Tuberc Scand Suppl 1960;48:1-119. 8. Warren NG, Body BA, Dalton HP. An improved reagent for mycobacterial nitrate reductase tests. J Clin Microbiol 1983;18:546-9. 103 I. Pigment Production (1) Principle Mycobacteria synthesize carotenoid pigments in various amounts and kinds, depending upon the species. Using these variations in pigment production, the mycobacteria may be categorized into three groups: photochromogens, scotochromogens, and nonphoto- chromogens. Pigment formation is best observed in cultures that have isolated colonies. Photochromogens are organisms that produce pigmented colonies only after exposure to light. These organisms usually are white, cream, or buff when grown in the dark and show no pigmentation unless exposed to light. Testing for photochromogenicity should be done on young, actively-metabolizing, well-aerated cultures (4). Temperature of testing should be standardized (3). Mycobacterium kansasii is a photochromogen that turns bright yellow after light exposure. In contrast, some strains of M. simiae demonstrate photochromogenicity only after prolonged periods for both light exposure and pigment development. Scotochromogens are organisms that produce pigmented colonies whether grown in the dark or the light. Mycobacterium gordcnae and M. scrofulaceum are examples. Pigment production may increase if cultures are exposed continuously (2 weeks or more) to light. Myco- bacterium szulgai demonstrates a somewhat unique characteristic, being scotochromogenic at 37°C, but photochromogenic when grown at 25°C. This observation, although not unique, is strongly suggestive of M. szulgai (2). For this reason, all pigmented cultures should be checked for photoactivated pigment at both 37°C and 25°C. Nonphotochromogens are organisms whose pigment production is not affected by light. Although commonly nonpigmented, occa- sional strains exhibit colors that range from pale pastels to deep orange. Pigment production should be studied on all slow growers that are nonpigmented when grown in the dark, and on pigmented cultures that have not been grown continuously in the dark. Method eo Media and Supplies Egg-base medium slants Black paper or aluminum foil Incubator, 35 to 37°C e Controls The test procedure provides its own control. 104 eo Procedure (1) (2) (3) Inoculate five tubes of egg-base medium with a broth cul- ture of the test organism diluted sufficiently to yield iso- lated colonies. Shield four tubes from the light by wrapping them with aluminum foil or black paper. (a) Incubate two shielded cultures at 25°C. (b) Incubate the other two shielded cultures and the sin- gle unshielded culture at 37°C. (c) Occasionally, other incubation temperatures may be appropriate (e.g., suspected M. marinum or M. xenopi). Proceed as follows when growth is seen on the unshielded tube: (a) Remove shields from one tube at each incubation temperature, examine for growth, and record any pig- mentation (color plate 16). The second shielded tube (in each incubator) is the control culture for photoactive pigment production and should not be exposed to light. (b) If the examined tube(s) is(are) not pigmented, expose it (them) to light for 3 to 5 hours (a 60-W tungsten bulb placed 20 to 25 cm from the culture, or the equivalent in a fluorescent lamp, is adequate). Be certain the caps of tubes are kept loose during light exposure. (c) If culture grown at 37°C is pigmented, carefully note pigment on culture grown at 25°C (M. szulgai may be photochromogenic at 25°C, scotochromogenic at 37°C). (d) Replace shield(s) after light exposure. (e) Reincubate all cultures at appropriate temperature and examine after 24, 48, and 72 hours. Note: Photoactive pigment may appear more slowly in cul- tures grown at 25°C than in those grown at 37°C. e Results and Interpretation Compare light-exposed culture(s) with culture(s) maintained in shielded tube(s). « A culture that is pigmented both in the dark and the light is a scotochromogen e A culture that is nonpigmented in the dark but becomes yellow-to-orange after light exposure is a photochromogen Mycobacterium szulgai is one species that may be scoto- chromogentic at 37°C but photochromogenic at 25°C A culture that produces no pigment (or only pale pastel colors), either in the dark or after light exposure, may be labeled a nonphotochromogen 105 e Although rapid growers are readily segregated by their growth rate, they, too, may exhibit all the pigment varia- tions described above COLOR PLATE 16A. Nonphotochromogen (culture on left grown in dark, culture on right exposed to light). COLOR PLATE 16B. Photochromogen (culture on left grown in dark, culture on right exposed to light). COLOR PLATE 16C. Scotochromogen (culture on left grown in dark, culture on right exposed to light). 106 REFERENCES 1. Runyon EH, Karlson AG, Kubica GP, et al. Mycobacterium. In: Lennette EH, Spaulding EH, Truant JP, eds. Manual of clinical microbiology, 2nd ed. Washington, D.C.: American Society for Microbiology, 1974. 2. Selva-Sutter EA, Silcox VA, David HL. Differential identification of Mycobacterium szulgai and other scotochromogenic mycobacteria. J Clin Microbiol 1976;3:414-20. 3. Tsukamura M. Relationship between photochromogenicity and test temperatures in mycobacteria. J Clin Microbiol 1981;14:225-6. 4. Wayne LG, Doubek JR. The role of air in the photochromogenic behavior of Mycobacterium kansasii. Am J Clin Pathol 1964;42:431-5. J. PYRAZINAMIDASE (1) Principle The deamidation of pyrazinamide (PZA) to pyrazinoic acid and ammonia is helpful in separating M. marinum (positive in 4 days) from M. kansasii (negative), and weakly niacin-positive strains of M. bovis from M. tuberculosis and the M. avium complex. Mycobacterium bovis is pyrazinamidase negative even at 7 days, whereas both M. tuberculosis and the M. avium complex are positive within 4 days. Method e Media and Supplies Dubos broth base Distilled water Pyrazinamide Pyruvic acid, sodium salt Agar Screwcap tubes, 16 x 125-mm Ferrous ammonium sulfate e Preparation Substrate medium Dissolve 6.5 g of Dubos broth base in 1 liter of distilled water. Add 0.1 g of pyrazinamide, 2.0 g of pyruvic acid, sodium salt, and 15.0 g of agar. Heat to dissolve the agar and dispense in 5-ml amounts in 16 x 125-mm screwcap tubes. Autoclave at 121°C for 15 minutes. Allow the medium to harden in an upright position to form a butt. When stored at 5°C, the medium keeps for several months. 107 1% Ferrous ammonium sulfate Prepare a 1% ferrous ammonium sulfate solution just before use. For convenience, 0.1 g amounts may be preweighed and stored in sterile 16 x 125-mm screwcap tubes. Add 10 ml of sterile distilled water when the solution is needed. Controls Use M. avium complex as a postive control. A tube of uninoculated medium and either M. bovis or M. kansasii serve as negative controls. A “color control standard’ (developed by W. R. Butler, Myco- bacteriology Branch, Center for Infectious Diseases, CDC) may be prepared in the same manner as the substrate medium; sim- ply replace the pyrazinamide with the same amount (0.1 g) of pyrazinoic acid (2-pyrazine-carboxylic acid, 99%, from Aldrich Chemical Co., Milwaukee, WI). Procedures (1) Inoculate the surface of two tubes of medium with a heavy loopful of growth from an actively growing culture (2 to 3 weeks old). The inoculum should be heavy enough to be visible. (2) Incubate cultures and controls at 37°C. (3) After 4 days, add 1.0 ml of freshly prepared 1% ferrous ammonium sulfate solution to each unknown culture, to the color control standard, and to one each of the positive and negative controls. (4) Leave tubes at room temperature for 30 minutes and then examine for a pink band in the agar medium. (5) Refrigerate negative tubes for an additional 4 hours to mini- mize growth of contaminants from nonsterile ferrous ammonium sulfate solution and examine the medium again for a pink band in the agar. The color is easiest to detect by examining the tube against a white background and using incident room light (color plate 17). (6) If the 4-day tube is negative or doubtful, repeat the test at 7 days using the second tube. (7) If the 4-day tube is positive, the second tube may be dis- carded without further incubation. Results and Interpretation A pink band, which forms in the subsurface agar and diffuses into the medium, indicates the enzymatic hydrolysis of PZA to free pyrazinoic acid. Positive = pink band in agar. Negative = no pink band in agar. 108 = ; Cc Cc —- + COLOR PLATE 17. Pyrazinamidase. REFERENCES 1. Wayne, LG. Simple pyrazinamidase and urease tests for routine identification of mycobacteria. Am Rev Respir Dis 1974;109:147-51. K. Sodium Chloride Tolerance (1) Principle Mycobacteria vary in their ability to grow in the presence of, or tolerate, 5% sodium chloride (NaCl). Most rapid growers and the slowly growing M. triviale will grow on the NaCl-containing medium (2, 3). Some strains of M. flavescens also may grow on 5% sodium chloride. The inability of M. chelonae subsp. chelonae to grow on 5% sodium chloride facilitates its distinction from other members of the M. fortuitum complex. Method e Media and Supplies NaCl Egg-base medium (ATS or L-J) Screwcap tubes, 18 x 150-mm Recent reports suggest using 7H-11 medium for the sodium chloride tolerance test. (Tsang AY. 82nd Annual Meeting Am Soc Microbiol 1982, C183.) e Preparations Prepare two bottles of egg-base medium according to stan- dard formulation, page 49. To one batch of 300 ml of complete egg medium, add 15 g of NaCl. Mix well. The second batch of egg medium is used as a growth control and contains no NaCl. Dispense each medium in 7- to 8-ml amounts into 18 x 150-mm screwcap tubes and inspissate in slanted position at 85°C for 50 minutes. 109 e Controls The test procedure provides its own control. e Procedure (1) Inoculate both a control slant and an NaCl-containing slant with 0.1 ml of barely turbid suspension of a broth culture (or a saline suspension from a fresh slant culture). (2) Incubate at 28°C. (3) Examine weekly for growth. (4) After 4 weeks’ incubation, make final reading and discard tubes. e Results and Interpretation If more than 50 colonies develop on the NaCl tube within the 4-week period, the test is considered positive. Inoculum on the control medium should yield numerous colonies in the same period of time. Positive = growth on both the NaCl and control tubes. Negative = growth only on control medium. REFERENCES 1. Kestle DG, Abbott VD, Kubica GP. Differential identification of mycobacteria. Il. Subgroups of groups Il and Ill (Runyon) with different clinical significance. Am Rev Respir Dis 1967;95:1041-52. 2. Kubica GP. Differential identification of mycobacteria. VII. Key features for identifica- tion of clinically significant mycobacteria. Am Rev Respir Dis 1973;107:9-21. 3. Silcox VA, Good RC, Floyd MM. Identification of clinically significant Mycobacterium fortuitum complex isolates. J Clin Microbiol 1981;14:686-91. L. Tellurite Reduction (1) Principle During peak growth in liquid culture, mycobacteria reduce potas- sium tellurite at variable rates. In the test, tellurite acts as an artificial electron acceptor and is reduced to black, metallic tellurium at sites of oxidation-reduction activity in the bacterial cells. Although mycobac- teria may be placed into one of several groups, depending upon their speed of tellurite reduction, the ability of the organisms to reduce tellurite in 3 days is most useful for separating M. avium complex (positive) from most other nonphotochromogens (negative). Most rapid growers also reduce tellurite in 3 days. 110 Method e Media and Supplies Middlebrook 7H-9 medium, commercial powdered base Tween 80 Albumin-dextrose-catalase (ADC) enrichment for 7H-9 medium, commercial Screwcap tubes, 20 x 150-mm or 16 x 125-mm and 13 x 100-mm Potassium tellurite, 0.2% e Preparation 7H-9 liquid medium Prepare according to directions 900 ml of Middlebrook 7H-9 liquid medium from the powdered base, and supplement with 0.5 ml Tween 80. Do not use glycerol. Autoclave at 121°C for 15 minutes, cool to 45°C, and aseptically add 100 ml of ADC enrichment. Aseptically dispense no more than 5 ml of com- pleted medium into 20 x 150-mm screwcap tubes (or no more than 2.5 ml into 16 x 125-mm screwcap tubes). 0.2% Potassium tellurite Dissolve 0.1 g of potassium tellurite in 50 ml of distilled water. Dispense in 2-ml amounts into 13 x 100-mm screwcap tubes and autoclave at 121°C for 10 minutes. Store in refrigerator. e Controls Use a tube inoculated with M. avium as a positive control. Tube inoculated with M. terrae complex serves as a negative control. e Procedure (1) Inoculate the 7H-9 liquid medium with a heavy spadeful of test organisms taken from a young, actively growing cul- ture slant. (2) Incubate at 35°C for 7 days only. If growth is not heavy at 7 days, reinoculate a new liquid culture with a heavy spade- ful of growth for possible retest the following week. The poorly growing culture may be tested at 7 days, but do not reincubate for additional time in the hope of getting heav- ier growth. All inoculated cultures should be hand shaken daily to encourage heavy growth by 7 days. (3) Add 2 drops of sterile potassium tellurite solution (1 drop if 2.5-ml amounts in 16 x 125-mm tubes were inoculated) to each test culture and to controls, and shake the tubes to mix. Discard any unused tellurite solution. (4) Reincubate all cultures at 35°C for 3 days. Do not shake tubes during this time. 111 (6) On third day, examine sedimented cells in each culture tube, taking care not to shake tubes (color plate 18). e Results and Interpretation Positive = formation of a black precipitate of metallic tellurium in and around the sedimented bacterial cells. Negative = growth of cells without a black precipitate. Some species produce a light brown or gray precipitate that should be recorded as a negative test. COLOR PLATE 18. Tellurite reduction test. e Precaution Remember, if after 7 days of incubation, cultures are not heav- ily turbid, repeat the test with a fresh, heavily turbid broth culture. This will minimize questionable or suspected false negative tests. REFERENCES 1. Kilburn JO, Silcox VA, Kubica GP. Differential identification of mycobacteria. V. The tellurite reduction test. Am Rev Respir Dis 1969;99:94-100. M. Thiophen-2-Carboxylic Acid Hydrazide (TCH) Suscep- tibility Test (1, 4) Principle This test is valuable for distinguishing M. bovis from M. tuberculo- sis (1, 2) and other nonchromogenic slowly growing mycobacteria (3, 4). Only M. bovis is susceptible to low concentrations of TCH, 1 to 5 pg/ml. Isoniazid-resistant strains of M. bovis may be resistant to TCH. Mycobacterium tuberculosis and other mycobacteria are usually resis- tant to the inhibitory action of this compound. 112 Method Media and Supplies Middlebrook 7H-10 medium Thiophen-2-carboxylic acid hydrazide (Aldrich Chemical Co., Milwaukee, WI) Sterile screwcap tubes or four-section petri plates (plastic quad- rant plates). Preparation Prepare two batches of complete, enriched Middlebrook 7H-10 medium. One batch is poured as a drug-free control medium; to the other is added sufficient filter-sterilized TCH to make a final concentration of 2 ug/ml. Dispense each medium into sterile, screwcap tubes or, preferably, into four-sectioned petri dishes. Controls The test provides its own control. eo Procedure REFERENCES (1) Dilute a 7-day-old liquid test culture to 10-3 and 10-5 (1:1000 and 1:100,000) in sterile saline or water. (2) Inoculate one control and one drug-containing medium with 0.1 ml of each dilution. (3) Incubate for 3 weeks at 35°C in an atmosphere of 10% car- bon dioxide - 90% air. Results and Interpretation Record the organisms as resistant to TCH if growth on the drug-containing medium is equal to or greater than 1% of that observed on the drug-free control medium. 1. Bonicke R. Die differenzierung humaner und boviner Tuberkelbakterien mit Hilfe von Thiopen-2-carbonséure-hydrazid. Naturwissenschaften 1958;46:392. 2. Harrington R, Karlson AG. Differentiation between M. tuberculosis and M. bovis by in-vitro procedures. Am J Vet Res 1966;27:1193-6. 3. Runyon EH, Karlson AG, Kubica GP, Wayne LG. Mycobacterium. In: Lennette EH, Spaulding EH, Truant JP, eds. Manual of clinical microbiology, 2nd ed. Washington, D.C.: American Society for Microbiology, 1974:148-74. 4. Vestal AL, Kubica GP. Differential identificaton of mycobacteria lll. Use of thiaceta- zone, thiophen-2-carboxylic acid hydrazide and triphenyltetrazolium chloride. Scand J Respir Dis 1967;48:142-8. 113 N. Tween Hydrolysis (2) Principle The enzymatic hydrolysis of Tween 80 (with a few exceptions) is used to separate the potentially pathogenic (negative) from the com- monly saprophytic (positive) species among the slowly growing scotochromogens and nonphotochromogens. The test medium con- tains Tween 80 and the indicator neutral red in a buffered neutral (pH 7) solution. Contrary to common impression, the color change in the Tween hydrolysis test is not due to a pH shift related to release of oleic acid from the hydrolytic cleavage of Tween 80. Normally, neu- tral red is red at pH 7, but when bound by lipids or by Tween 80, the indicator dye takes on the amber or straw color that it commonly has at more alkaline pH. When (and if) the bacterial esterases split the Tween 80, it no longer acts to bind the indicator dye and the neutral red then reverts to its usual color at pH 7, which is red, and the test is called positive. Method e Media and supplies Tween 80 0.067 M phosphate buffer, pH 7.0 Aqueous neutral red, 0.1% Screwcap tubes, 16 x 125 mm e Preparation Phosphate buffer, pH 7.0 Prepare stock solutions as follows: a. Disodium phosphate, 0.067 M. Dissolve 9.47 g of anhydrous Na,HPO, in one liter of dis- tilled water. b. Monopotassium phosphate, 0.067 M. Dissolve 9.07 g of KH,PO,4 in one liter of distilled water. To prepare 100 ml of pH 7.0 buffer, mix 61.1 ml of solution “a” with 38.9 ml of solution “b"’. Check the pH of the final solution. Substrate medium Add the following, in order, to 100 ml of pH 7.0 phosphate buffer: 0.5 ml of Tween 80 and 2 ml of a 0.1% aqueous solution of neutral red. Dispense the substrate in 2-m| amounts into 16 x 125-mm screwcap tubes. Autoclave at 121°C for 10 minutes. The substrate should be straw or amber color after autoclaving. Store in the dark at 5°C for no more than 2 weeks. Note: A commercially available concentrated TB Tween hydrol- ysis reagent is stable for up to 1 year and provides excellent test results (1). Follow manufacturer's directions. 114 e Controls Use an uninoculated tube of substrate for the negative color control. A tube inoculated with M. kansasii serves as a positive control. A tube inoculated wth M. bovis (BCG) or M. avium complex may be used as a negative control. e Procedure (1) Inoculate the substrate with a spadeful of test organisms from an actively growing, young culture slant. (2) Incubate at 35°C. (3) Examine after 1, 5, and 10 days. Do not shake tubes while reading them. e Results and Interpretation Positive = substrate turns pink to red. Record the number of days required for the first appearance of a pink-to-red color (color plate 19). Negative = substrate remains amber-colored after 10 days’ incubation. Positive reactors should be recorded for up to 10 days, at which time remaining negative reactors should be recorded as such and discarded. COLOR PLATE 19. Tween hydrolysis. eo Precautions Occasionally the sedimented cells will be red in an amber- colored fluid. This is a positive neutral red test, but a negative Tween hydrolysis test. A positive hydrolysis test is recorded only when the entire liquid medium has turned salmon pink to red. Sometimes there is a bleaching of the supernatant fluid dur- ing incubation. These tests should be repeated and recorded as negative if the bleaching recurs. 115 REFERENCES 1. Kilburn JO, O'Donnell KF, Silcox VA, David HL. Preparation of a stable mycobacte- rial Tween hydrolysis test substrate. Appl Microbiol 1973; 26:826. 2. Wayne LG, Doubek JR, Russell RL. Classification and identification of mycobacteria. |. Tests employing Tween 80 as substrate. Am Rev Respir Dis 1964;90:588-97. O. Tween Opacity (1) Principle The test is especially helpful in separating M. flavescens (positive in 1 week) from other pigmented slow growers. The exact mechanism of the reaction is not known, but Wayne et al. believe it is due to oleic acid or oleate salts released from the Tween 80. Method e Media and Supplies Dubos oleic acid agar base Tween 80 Distilled water Dubos albumin enrichment Screwcap tubes, 16 x 125-mm e Preparations Test medium Add 4 g of commercial Dubos oleic agar base and 5 ml of Tween 80 to 180 ml of distilled water. Autoclave at 121°C for 10 minutes. Cool to 56°C in a water bath before adding 20 ml of commercial Dubos medium albumin. Dispense 5-ml amounts into 16 x 125-mm screwcap tubes. Allow medium to solidify with tubes in an upright position. Control medium Prepare medium in identical manner as for the test medium, except add only 0.04 ml of Tween 80 (or 0.4 ml of a 10% aqueous solution of Tween 80). e Controls The test provides its own control. e Procedure (1) Grow test cultures in 7H-9 broth for 7 days. (2) Dilute broth culture 1:100 in sterile saline or water. (3) Inoculate the surface of both a control and test medium with 0.1 ml of the diluted suspension. 116 (4) Incubate tubes at 35°C. (5) Examine weekly for 6 weeks. e Results and Interpretation Positive = formation of an opaque ring below the surface of the test medium. Record results when opacity is first seen because the ring will become more diffuse with time. The agar just above the opaque ring tends to become clear (i.e., that portion just below the sur- face of the medium). REFERENCES 1. Wayne LG, Doubek JR, Russell RL. Classification and identification of mycobacteria. |. Tests employing Tween 80 as substrate. Am Rev Respir Dis 1964;90:588-97. P. Urease (3, 4) Principle The ability of a culture to hydrolyze urea (releasing ammonia) is useful in identifying both scotochromogens and nonphotochromogens (3, 4). M. scrofulaceum, M. szulgai, M. flavescens, M. bovis, M. tuberculosis, and M. gastri are positive, whereas M. avium complex, M. xenopi, M. terrae complex, and M. gordonae are negative. Methods 1. Murphy-Hawkins Disk Method (1) eo Media and Supplies Difco urea differentiation disks Sterile water Screwcap tubes, 13 x 100-mm e Preparation Place a single urea differentiation disk into a sterile 13 x 100-mm screwcap tube containing 0.5 ml of sterile water. e Controls Use M. scrofulaceum or M. fortuitum as a positive control. An uninoculated substrate tube and a tube inoculated with urease-negative M. gordonae serve as negative controls. 117 2. e Procedure (1) Into a substrate tube, emulsify a heavy spadeful of test organisms from a 3-week-old egg medium cul- ture slant. (2) Incubate tubes at 35°C. (3) Read results at 1 hour and daily thereafter for 3 days. Results and Interpretation Positive = substrate color change to a moderate-to-deep red (cerise). Negative = no color change or only a pale pink (color plate 20). COLOR PLATE 20. Urease - Murphy-Hawkins Disk Method. e Precautions Occasional test difficulties may be attributed to the fail- ure to carefully acid-wash and thoroughly rinse the test tubes used. Also, the possibility of encountering a bad batch of paper disks should be considered. Careful attention to the reaction of the control tubes will indicate when test reactions are amiss. Steadham Method: Texas Urease Test (2) eo Media and Supplies Peptone Dextrose Sodium chloride (NaCl) Monopotassium phosphate (KH,PO,) Urea Phenol red, 1% Tween 80 Distilled water 118 e Preparation Dissolve the following ingredients in 1 liter of distilled water: 1 g of peptone, 1 g of dextrose, 5 g of NaCl, 0.4 g of KH,PO,4, 20 g of urea, 1 ml of a 1% solution of phenol red sodium salt, 0.1 ml of Tween 80 (or 1 ml of 10% solution of Tween 80). Adjust final pH to 5.8 = 0.1 with NaOH. Sterilize by membrane filtration through 0.22 pm pore-size filters. Aseptically dispense 1.5-ml amounts into 18 x 125-mm sterile, screwcap tubes. Tighten caps and store at 5°C for up to 2 months. e Control Same as mentioned in previous urease methods. e Procedure (1) Macerate a spadeful of growth from a young, actively growing slant into the urea broth. (2) Incubate at 35°C without CO,. (3) Read test at 1, 3, and 7 days. e Results and Interpretation Positive = color change in broth from yellow to dark pink or red (color plate 21). Negative = no color change. A light pink is recorded as negative or doubtful and should be repeated. Steadham has recommended the use of nitrate reduction color standards (see page 102) to better stan- dardize color interpretation. Hi COLOR PLATE 21. Urease - Steadham Method. Wayne Method (5) eo Media and Supplies Difco-Bacto concentrate urea agar Sterile distilled water Screwcap tubes, 13 x 100-mm 119 e Preparation Aseptically mix 1 part of Difco-Bacto urea agar concen- trate with 9 parts of sterile distilled water. Do not add agar. Dispense 3-ml amounts into 13 x 100-mm sterile screwcap tubes. e Controls Use substrate inoculated with M. scrofulaceum or M. bovis as a positive control. An uninoculated tube of sub- strate and a tube inoculated with urease-negative M. gordonae serve as negative controls. e Procedures (1) Inoculate the liquid substrate with a spadeful of growth from a young actively growing culture slant. (2) Incubate at 35°C. (3) Observe for a color change (color plate 22). e Results and Interpretation Positive = color change from amber to pink or red. Negative = no color change. COLOR PLATE 22. Urease - Wayne Method. REFERENCES 1. Murphy DB, Hawkins JE. Use of urease test disks in the identification of mycobacteria. J Clin Microbiol 1975;1:465-8. 2. Steadham JE. Reliable urease test for identification of mycobacteria. J Clin Microbiol 1979;10:134-137. 3. Toda T, Hagihara Y, Takeya K. A simple urease test for the classification of mycobacteria. Am Rev Respir Dis 1960;83:775-61. 4. Urabe K, Saito H. The urease activity of mycobacteria. Am Rev Respir Dis 1964;90: 266-7. 5. Wayne LG. Simple pyrazinamidase and urease tests for routine identification of mycobacteria. Am Rev Respir Dis 1974;109:147-51. 120 Culture Examination and Identification Culture Examination and Identification In 1983, there were 54 recognized species of mycobacteria (11). One of these (Mycobacterium leprae) does not grow on ordinary culture media and six are pathogens of animals (M. farcinogenes, M. lepraemurium, M. microti, M. paratuberculosis, M. porcinum, M. senegalense). An additional 23 species of rapidly growing mycobac- teria rarely infect humans, are commonly saprophytic, and usually are lumped together as “other rapid growers.” Only if such isolates are regarded as potentially pathogenic is any effort made to identify them. A few slowly growing mycobacteria (M. africanum, M. asiati- cum, M. bovis, M. haemophilum, M. shimoidei, M. ulcerans) are seen so rarely in the United States that most laboratorians know them only by name. Two reasons for our lack of familiarity with some of these species may be their relatively recent appearance on the microbio- logic scene and their currently limited geographic distribution. But the frequency of international travel and the emigration of people to other countries could change this picture in the future. Thus, there are just over 20 species of mycobacteria, isolated with variable frequencies, about which most mycobacteriologists should be aware. In the 1950s, before they had names, the so-called “atypical” mycobacteria were subdivided by Runyon (4-6,9) into four groups on the basis of pigment production and growth rate. Although mycobac- teria should be precisely identified by species name, occasional recent isolates or unknown organisms may be referred to by vernacular names—photochromogens, scotochromogens, nonphotochromogens, and rapid growers—to facilitate discussion about and ultimate identi- fication of these organisms. Too, the use of complexes (3,7,8,10)— e.g., M. avium complex, M. fortuitum complex, M. terrae complex— permits the lumping together of species of common medical impor- tance, when more precise characterization seems to be of little clini- cal value. There is some overlap of the original pigment and growth rate groups of Runyon (4), but the initial subdivision of mycobacteria based upon pigmentation and speed of growth is indeed a helpful tool for identification (figure 17). Photochromogens were originally defined as mycobacteria that were nonpigmented when grown in the dark but acquired a lemon-yellow color after exposure either to daylight or artificial light (incandescent 121 or fluorescent) (5). All species currently in this group are potentially pathogenic: M. asiaticum, M. kansasii, M. marinum, and M. simiae. Of 368 strains in this group (figure 17), 89% were photochromogenic, 2% were “‘albino’’ mutants that produced no pigment, and 0.4% were scotochromogenic mutants that were yellow to orange regardless of light exposure. Nine percent of the isolates grew rapidly: 80% of these were identified as M. marinum, a species that can grow rapidly if inoculated heavily or if grown at its optimum temperature of 32 to 33°C; 20% were identified as M. kansasii which only rarely will grow rapidly, the most common reason being the use of an unusually heavy inoculum. The word Scotochromogen is derived from the Greek, skotos, mean- ing “in the dark’’; hence, an organism pigmented both in dark and light (independent of light exposure, although the latter may intensify the color). Species in this group include both potential pathogens (M. scrofulaceum, M. szulgai, and M. xenopi/, the last two exhibiting strange pigment features) and common saprophytes (M. gordonae and M. flavescens, the latter often grouped with rapid growers). Of 707 strains, 94.5% were truly scotochromogenic, 1.4% photochromo- genic (some undoubtedly M. szulgai), 0.5% were nonphotochromo- genic (although speciated as a known ““scotochrome’’), and 3.5% were rapid growers (75% of these were M. flavescens, an organism of intermediate growth rate; 15% were M. scrofulaceum and 10% M. gordonae; the latter two probably “misplaced” by use of a heavy inoculum). Although most Nonphotochromogens are indeed nonpigmented (figure 17), the name does suggest that some may be pigmented but that the pigment is not affected by light. Eighty-seven percent of 1528 FIGURE 17. Separation of mycobacteria on the sole basis of growth rate and pigment production Percent reacting ast Identified” as Photo- Scotochrome Non Photo- Rapid (No. of strains) chrome chrome Grower Photochrome (368) 89 0.4 2.0 9.0 Scotochrome (707) 1.4 94.5 0.5 3.5 Nonphotochrome (1528) 1.7 8.3 87 2.9 Rapid Grower (620) 2.9 97.1 1''Percentage reacting as’ determined only by pigmentation and growth rate. *Precise identification determined by reaction patterns in a battery of biochemical tests. 122 isolates were nonchromogenic, 1.7% exhibited photochromogenic pigment, and 8.3% were scotochromogenic, producing bright yellow- to-orange pigment. Nearly 3% of the nonphotochromogens grew rapidly; 82% of these were identified as M. terrae, 9% as M. gastri, about 2.5% each as M. avium and M. triviale (the last species once referred to as a ‘‘slow-growing rapid grower”), and 4% as M. tubercu- losis. Perhaps more than anything, this last observation points up the need to be especially careful about the inoculum size when preparing subcultures to determine growth rate. Although most (97.1%) Rapid Growers are identified by their ability to yield fully mature, grossly visible colonies in less than 7 days, 2.9% of 620 strains examined grew slowly (for whatever reason) and were nonpigmented. A number of strains of M. chelonae may grow slowly when incubated at 35 to 37°C, but when incubated at their optimum temperature of 28°C, they mature in less than 1 week. It is reassuring that these subgroups of mycobacteria, effected on the basis of pigment and growth rate, remain valid today even in the face of our more refined and precise taxonomic classification. Such a preliminary subgrouping often facilitates the precise identification of an isolated strain of Mycobacterium by directing the selection of the most appropriate differential taxonomic tests. Annual reviews of the numbers and species of mycobacteria recov- ered by the State health department laboratories of the United States have been conducted by CDC since 1979; two of these reviews have been published (1,2). Since 1980, these studies included both patho- genic and saprophytic species of mycobacteria; table 3 shows a provi- sional summary of the total number of each mycobacterial species that has been reported annually by the State laboratories from 1979 to 1981. By using this information, together with test probability data from the Mycobacteriology Branch (summarized in table 4 for the most commonly encountered species), it is possible to calculate the reliability of a specific identification of a given mycobacterial species. Additionally, the probability data in table 4 also enable the theoretical determination of how frequently strains of other species may present the same differential test pattern as the species being studied; i.e., how often may other species appear as “look alikes,"” and thus pres- ent the potential of a misidentification? If one knows the number of patient isolates of each species of Mycobacterium recovered in the State health department laboratories (table 3) and the reliabilities of positive or negative reactions in the various in vitro tests used for differential identification of mycobacteria (table 4), it is possible to determine the statistical probabilities of (a) a correct identification of a given “‘subject species,” and (b) the likeli- hood that strains of a different species may be confused with the “subject species.” The methods and programs used to calculate the 123 TABLE 3. Numbers of indicated species of Mycobacterium recovered by state health departments of the United States, 1979-1981* 1979 1980 1981 Mycobacterium Species Number Ave. no./| Number Ave. no./ Number Ave. no./ GP (% of total) SHD (% of total) SHD (% of total) SHD 2 | asiaticum 2(0.007) <1 4(0.01) <1 12(0.03) <1 8 | kansasii 799(2.90) 16| 1133(2.76) 23 902(2.32) 18 § marinum 75(0.27) 2| 162(0.39) 3| 108(0.28) 2 S| simiae 39(0.14) 1 7800.19) 2| 2900.07) 1 2 | others 2(0.007) <1 4(0.01) <1 22(0.06) <1 ®| subtotal 917(3.33) 21 1381(3.36) | 30 1073(2.77) | 25 flavescens 26(0.09) 1| 338(0.82) 7| 335(0.86) 7 i gordonae 911(3.31) 18| 6008(14.61) 120| 6735(17.32) 135 8 | serofulaceum 740(2.69) 15| 763(1.86) 15| 889(2.29) 18 2 szulgai 45(0.16) 1 48(0.12) 1 49(0.13) 1 8 | xenopi 51(0.19) 1 85(0.21) 2 94(0.24) 2 3 Others 911(3.31) 18] 412(1.00) 8| 360(0.93) 7 Subtotal 2684(9.74) [54 7654(18.62) | 153 8462(21.84) | 170 avium 4462(16.19) 89 6980(16.98) 140| 7424(19.09) 149 @ | gastri 106(0.26) 2 121(0.31) 2 & haemophilum E | maimoense 2(0.007) <1 12(0.03) <1 21(0.05) <1 £ shimoidei o | terrae cpix 180(0.65) 4 867(2.11) 17 777(2.00) 16 a triviale 2 | others 665(2.41) 13] 540(1.31) 11 532(1.37) 1 Subtotal 5309(19.26) [107 8505(20.69) | 171 8875(22.91) | 179 africanum x | bovis 10(0.04) <1 27(0.07) 1 20(0.05) <1 S| sca 1(0.0004) <1 1(0.002) <1 8(0.02) <1 @ | tuberculosis 17015(61.70) 340[20980(51.03) 420[17541(45.10) 351 Subtotal 17026(61.77) |342 21008(51.10)| 422 17569(45.34) | 353 g | chelonae 373(1.35) 8| 539(1.31) 11] 600(1.54) 12 2 | fortuitum 1099(3.99) 22| 1054(2.56) 21] 1248(3.21) 25 & | fortuitum cplix 62(0.23) 1] 530(1.29) 11 518(1.33) 10 2 | others 93(0.34) 2| 420(1.02) 8| 383(0.99) 8 @ | Subtotal 1627(5.90) 33 2543(6.19) | 51 2749(7.09) | 55 Unknown 1(0.004) <1 21(0.05) <1 18(0.05) <1 Grand Totals 27564(100) ~558( 41112(100) ~828| 38746(100) ~782 *Provisional data; late or corrected data not included. 124 TABLE 4. Identification of clinically important mycobacteria” © > NE: °c Arylsulfatase Carbon Sources Catalase 2 g Sle So cd c aE SPECIES ,o | 2|zaz|.8%8 Semi. | Heat 5% S| |5. | .5] 6 %» ” 3° i ts [aSy|3ca|scs| 3 2 Sodium| Inosi- [Manni ’ > £5 c sg | cg [£8 £5 £ Z8 zZ3 2 (subspecies) sg |gz2|3Th|E2e ) Quant [Stable x $3 g sS5( 52352 5% 3 go0 Sty 2 a cS [252 " 1] >25| Day Wk Citrate| tol tol “45 (68°C) SE 8% 8 £3 3 £ 3 =o 3x [8 so | 382 3 os |RP6c|0xn|fES > £5 | 6s z za | 25 |6& ee | RET | 28] 5 M. fortuitum biovar (87) 3 _ _ _ fortuitum EARN 22-40 , J (99) 99) | (99) 45 + + + — + + + + +/— + rrr om: : 07) | { 199) F— ~ = om [S41 [Loo [Leer (09) |{ 09) 75) [L971 | (99) |{ 150) 93) Blo Smooth | 22-37 N (99) (99) (99) # M. fortuitum = + + 3rd. biovariant complex Smooth | 22:37 (99) (99) (99) 2 [ M chelonae R (99) — 9 g subsp. chelonae } 22:35 (97) 09) | (09) | (99) = (99) ‘ s M. chelonae Sreoats > + 4 - — = >45 —/* 98) + — — + + + —/+ (91) + 5 subsp. abscessus Rough (22.40 (96) (98) | (99) | (99) | (99) 197) (53/47) 96) |( (92) |[( (99) 90) [( 85) (198) |(46/54) (99) a M. chelonae 60/40, + — = + - « (turtle-like) 22:30 (99) (99) (99) (Tan) (99) (99) R(307) > = - = oy = ” = + = + — M. fallax (Looks ike M. tb.) Rough | 30-37 | 537°) | N=9%% | (99) 99) | (99) | (99) (74) | (99) | (99) | (99) | (99) (99) 99) | (99) (96) Smooth 2 po. — + — + + + + -— + — et — + + + + Other Rapid Growers an 17:52 |) rd (72) (75) (70) (59) (60) (66) (63) (56) (52) (95) (51) (52) (87) (99) (75) V (66) M. ulcerans Soa 28-33 Ss N — = + — = — + a - | Rou i — = — rp = + + + -— —_— + + — + i M. tuberculosis hay 33-39 S N=99%| (99) (93) (99) (99) (99) (98) (99) (98) (99) (70) (92) (68) (83) (98) E = bd = er — + 3S M. africanum Rough | 35-38 S N (99) —- - (99) (99) - = - (99) (99) Var oe (99) jo M bovi THIO=R — — — — — == os — ow = so —_ -_ + . bovis LJ=S 35-38 S N=99%| (99) (87) (97) (92) (99) (95) (94) (98) (99) (55) (94) (84) (75) (99) No “X" Colony | Smooth N 85% — + i + —~ — — + y — + + — = — M. avium complex (99) Rough | 22.45 |S=99% | 5 12% (99) (51) (99) (76) (68) (99) 92) [Fe 99] (99) (82) (99) (98) (99) (99) ; (99) N 80% + + — + w= — ¥ mm — + = == “ M. xenopi lsmoom x | 35-45 S S 20% (76) (99) 9) (83) (99) (99) (94) (51) (99) (59) (99) (99) (99) c # M. shimoidei Rough | 30-45 s N — - - + — — + _ + 3 _ 5 : S=80% — + — = wr — — Fa == ~ : oe = £| Mgastri No | |smoom| 25.40 |R-20% | N=99%| (gg) (99) 99) | (99) ©4) | 99) | 99) |@2 63] 69) | 7a) | 00) | 99) | 09) | 179) 8 XC Smooth S=77% | N 95% —_— —_ + + —— — — + © (88) wo — + + — = 2 M. terrae complex Colony Alam 22-37 | R=23% te (99) (51) (99) (96) (99) (77) (99) (72) [aa 74 (94) (75) (99) (99) (93) (91) 5 I” oo = + + : - = + leo so] 4 — + + = M. triviale (99) Rough | 22-37 S N=99%| (66) (99) (93) (99) (99) (97) (99) Jag 7a | (99) (99) (99) (99) (86) 2 s N=90% = = = + = - + = + + + cs M. malmoense smooth | 22-37 S=10%] (99) (89) (99) (69) (99) (99) 99 1 gu (99) (97) (67) M. haemophilum (Needs hemin) | Rough | 25-35 S N - = - - BD - * - - - or P=91% — — + + wy + ms . — + + — + M. simiae Smooth | 22-37 S N= 9%[ (99) (84) (39) (95) (99) (85) (85) (99) (85) (99) (85) (64) 2 Ji S-98% | £.29% — + + + — =~ + — + ~ = + + — + £ M. kansasii Sa | 25:40 [R=2% | 25% | 09) | 8) 99) | (95) 99) | (99) | (99) [226] (99) | (8a) | (99) | (99) | (64) | (97) c . R=61% 2 + = = Er 2 = - + — + + + § M. marinum smooth| 25-35 |5=399% | P=99%( (66) (99) (79) (58) (99) (99) (80) (96) (99) (97) (79) (99) (99) (99) s ° — — + + = - — — + + — £ M. asiaticum smooth| 33-37 |S=99 P=99%[ (99) (67) 91) (91) (99) (99) (99) (70) (99) (91) (99) Smooth S-95% | S=97%| — — + + — — — 1 — + = — ¥ 2 M. scrofulaceum Rough | 22-37 |R-5y, | p=3% | (99) (64) (93) (96) (99) (99) (85) | (99) (99) (57) (99) (98) (99) (99) ¥ ) No Smooth 5 a _ si + + J + — + + + = 1 E | M szulgai axe )| Rovan| 2237 | S|, | (99) | (62) 99) | 81) 99) [ (99) (99) | (56) [ (99) | (50) | (99) | (94) © = c V S=99% i it + + + -— — Po - + — — + + a _- % | M gordonae 2 MN smoom| 22:37 | R=1% | 799% 99) | (67) ©2) | (97) 99) | (99) | (95) [1B] (99) | (77) | 98) | (99) | (99) | (85) 2 © R=59% | 5-99 = + + + — — + + + — + + + + a | M. flavescens smoot | 25:42 15-419, °| (84) | (78) ®5) | (99) 99) | (99) | (96) | (99) | (71) | 67) | (99) | (97) | (99) | (71) 1 Test reactions listed as “+ or "=", tollowed by percentage of strains reacting as indicated. If no percentage is given, or space is blank, insufficient data were available or test is of no apparent value. 2 Pyrazinamidase data (Wayne Method) from both Wayne and Hawkins. Unless indicated, results are those at 4 days. Strains of M. tuberculosis resistant to PZA are often pyrazinamidase negative. Within M. terrae complex, M. nonchromogenicum usually +" and M. terrae usually '— 3 Urease data (Murphy-Hawkins Disk Method) primarily from Dr. Jean E. Hawkins. » Data on M. fallax from Levy-Frebault et a/ (1.J.S.B. 33:336, 1983). 5 Several strains in this complex have varied in carbon sources as follows: ‘+’ sodium citrate, "+" mannitol, "'— 125 inositol. probabilities are beyond the scope of this brief chapter. Rather, for each species within the four pigment-growth-rate subgroups of mycobacteria, we have listed (a) the key tests to identify the species; (b) the probability that any strain of the species will give this exact differential pattern; (c) a list of other species that may be seen often enough in your laboratory that they have the mathematical possibil- ity of looking like the “subject species’ you are trying to identify; and (d) a list of additional tests that may be used to facilitate the precise separation of ‘‘subject species’ from “look-alike” species (see later, tables 5B to 9B). Mycobacterium tuberculosis is still the most important cultivable acid-fast organism isolated from humans (table 3), representing over 50% of patient isolates received by State health department labora- tories of the United States in 1979 and 1980. Provisional data for 1981 list M. tuberculosis as 45% of all isolates. The next most commonly encountered group is the nonphotochro- mogens, accounting collectively for 21% or more of all acid-fast bacilli. Within this group, M. avium and M. terrae complex together account for 92% of all nonphotochromogenic mycobacteria seen (exclusive of M. tuberculosis). The scotochromogens represented 19% of total acid-fast isolates in 1980 and 22% in 1981; the most common species of the group is M. gordonae (15% to 17% of all isolates). The generally greater suscepti- bility of M. gordonae to digestion-decontamination procedures used for primary isolation of mycobacteria makes this an ideal “indicator organism’’ for digestant toxicity; if this commonly saprophytic myco- bacterium is not recovered from about 5% of clinical specimens, it is possible the laboratory (a) is using a very toxic digestion-decontami- nation procedure, (b) is not generating sufficient g-forces to ade- quately sediment acia-fast bacilli from treated specimens, (c) is suffer- ing from a combination of both factors, or (d) is not located in an area where M. gordonae is a common environmental contaminant. Among the scotochromogenic mycobacteria, the species M. gordonae and M. scrofulaceum account for 88% (1980) to 90% (1981) of all scoto- chromogens. Of the remaining two groups, rapid growers accounted for 6% to 7% of all isolates, with members of the M. fortuitum complex account- ing for 83% to 86% of the rapidly growing species. The photochro- mogens provide just over 3% of the total acid-fast bacilli recovered, and M. kansasii and M. marinum together account for 83% to 94% of this total. Many laboratory workers find it helpful to place an unknown acid- fast isolate into one of the four pigment-growth-rate subgroups before deciding what “key tests’’ will best speciate the unknown. As shown in figure 17, a small percentage of isolates from each of the four 127 pigment-growth-rate groups does not produce the expected pigment or growth-rate charactersitics, but for the most part, initial placement of these organisms into one of these groups does facilitate identifica- tion within each subgroup. For the reference laboratory, we recommend that an unknown myco- bacterium be identified by using a young, actively metabolizing sub- culture of the organism. This young subculture should be used to inoculate all, or most, of the differential test media listed in the diag- nostic key (table 4), including “growth rate’’ media inoculated with bacterial suspensions diluted sufficiently that the final, mature growth yields well-isolated colonies; only in this way is it possible to get a true determination of growth rate (i.e., < or >7 days), as shown in figure 18. While growth rate and pigment production are being assessed (figure 18), the other differential tests (inoculated at the same time) are incubating and being reported. In the final analysis, some of these differential tests may not be needed for the final identi- fication of the species in question, but the data should be recorded for each strain because (a) it will contribute to overall knowledge of test reliability and (b) if a current species is ultimately shown to be a mixture of two or more species or subspecies, the ready availability of this old recorded test data may facilitate subdivision and recogni- tion of the new “subgroups.” Once the growth rate and pigment subgroups are established, as in figure 18, the laboratorian may concentrate on those tests (already inoculated, now incubating, or in some cases, results already reported) most likely to yield the desired differential identification. Unknown Speed of Growth Subculture | 270 1 Slow (>7Days) Rapid (<7 Days) Pighent | 1 Nonchromogenic Chromogenic 1 Niacin 1 T Se ———— Positive Negative — Pigment Photoactive I I Ld = No Effect Yes No Nonphotochromogen Photochromogen Scotochromogen FIGURE 18. Separation by growth rate and pigment 128 A. Rapid Growers The major group of mycobacteria to be recognized first in the fore- going scheme is the Rapid Growers. The key differential tests for this group are listed in table 5A. In most cases, it is sufficient to separate the two potential pathogens, M. fortuitum and M. chelonae, from the remaining, commonly saprophytic ‘other rapid growers.” Both spe- cies may exhibit either smooth or rough colonies. Table 5B indicates that the key tests for identification of M. fortuitum are positive reac- tions in tests for 3-day arylsulfatase, MacConkey agar, nitrate reduc- tion, and iron uptake. Reference to table 4 indicates that the percent- age of strains of M. fortuitum that react positively in the preceding four tests is, respectively, 97%, 96%, 99%, and 99%. If these four percentages, converted to probabilities by first moving the decimal point two places to the left, are multiplied together (i.e., 0.97 x 0.96 x 0.99 x 0.99), they yield 0.913; this means that, theoretically, 91.3%, or 91 times out of 100, a given strain of M. fortuitum will react exactly as indicated in table 5B in the four differential tests. In the case of M. chelonae, the key reactions are positive in tests for 3-day arylsulfatase and growth on MacConkey agar, and negative reactions in tests for both nitrate reduction and iron uptake. The prob- ability of a correct diagnosis is 89%. See color plates 36 & 37, pages 156 & 157. TABLE 5A. Rapid growers M. fortuitum Complex ET cE |e] 05 | od 8 SS|SE|53| 8883 8% 5 |82 TX ET | S22 Se 8 So Ss Oo ca SS|S5|Sc| SS |8¢|S5| 3 |5¢ SS|EP |S | 22283 2R| 8 |°5 =e | 8 8 Ls $8 |S ISL 3 Test S Pigment _— RY —_— — — — = 4 /—1 3-Day Arylsulfatase + + + + + + + |=! MacConkey at 28°C + + + + + + + |r? NaCl Tolerance at 28°C + + + — + — a vt Nitrate Reduction + + + rn — — — vt Iron Uptake at 28°C + + + _— — | (Tan) [(Tan) | V! Utilize Citrate at 28°C — — + _— + + vt Utilize Mannitol at 28°C] — + + — — _— + | += Utilize Inositol at 28°C _— —_ + =o: os fs = vt > MCLO-M, chelonae-like-organism show <45 semiquant catalase, negative 68° catalase. V = Variable; +/— indicates most species positive, a few negative; —/ + indicates most species negative, a few positive. 129 For both these potentially pathogenic rapid growers, our studies indicate that most laboratories in the United States would not receive a sufficient number of strains of any other species of Mycobacterium for there to be a statistical probability that these other species might present a “look-alike’’ organism possessing the precise definitive fea- tures of M. fortuitum or M. chelonae. Occasional isolates of the “other” rapid growers might behave as members of the M. fortuitum com- plex in the key test patterns indicated in table 5B. Separation of these other species may be facilitated by positive reactions for (a) pigment production or (b) utilization of more than one substrate as a carbon source (citrate, mannitol, and inositol). The ability of an unknown rapid grower to utilize none, or only one, of the foregoing substrates is sometimes used by reference laboratories to identify to subspecies or to designate biovars of M. fortuitum and M. chelonae (table 5A). The mislabeling of one of the “saprophytic rapid growers’ as an M. fortuitum complex (pathogen) is not nearly as bad as is the reverse, i.e., labeling a pathogen as a saprophyte. In the former situation, both clinician and microbiologist are forced to confirm the suspect diagno- sis by repeated isolation, usually of large numbers of colonies, in the presence of pathologic change and in the absence of other possible pathogens. TABLE 5B. Problems in differential identification—rapid growers Subject Species, Possible Additional Key Test Pattern and “Look-Alike"” Differential (Probability of Correct ID) Species Tests? M. fortuitum 3-day Arylsulfatase MacConkey agar Nitrate reduction Iron uptake (0.91) M. chelonae None + + + + ? Other Rapid Growers ? | Carbon sources 3-day Arylsulfatase MacConkey agar + None Nitrate reduction — Iron uptake —- ? Other Rapid Growers ? | Carbon sources (0.89) 1 “Look-alike” species arranged in order of decreasing likelihood of confusion with “Subject species’. If species’ name is followed by asterisk (*), look-alike species may still be confused with subject species even after use of additional differential tests. 2 The “additional tests’ are commonly arranged in order of decreasing differential value. Those enclosed in parentheses are (a) available only in a few reference laboratories, (b) not done routinely or (c) in a few cases of limited value. 130 In addition to M. fortuitum and M. chelonae, there are 23 other species of rapid growers that may be recovered in the mycobacteri- ology laboratory from human or environmental sources. They are: M. agri M. neoaurum M. aichiense M. obuense M. aurum M. parafortuitum M. austroafricanum M. phlei M. chitae M. pulveris M. chubuense M. rhodesiae M. diernhoferi M. smegmatis M. duvalii M. sphagni M. fallax M. thermoresistibile M. gadium M. tokaiense M. gilvum M. vaccae M. komossense None of these 23 is commonly related to human disease and all are commonly lumped together as “other rapid growers,” with no fur- ther distinction attempted. One of the group, M. fallax, is listed sepa- rately in table 4, not because of its potential pathogenicity but because of its outward similarity to M. tuberculosis. Although M. fallax will grow rapidly (<5 days) when incubated at 30°C, incubation at 37°C results in much slower growth (12—21 days). Colonies are rough, eugonic, nonpigmented, and corded. It is also positive in nitrate reduc- tion and negative in 68°C catalase tests. But rapid growth at 30°C and a negative niacin test help to distinguish M. fallax from M. tuberculo- sis. Only time will tell the importance of this new species to clinical mycobacteriology. Niacin-Positive Nonchromogens Because M. tuberculosis is the species of Mycobacterium most com- monly encountered in the mycobacteriology laboratory (table 3), it is common to use the niacin test to segregate M. tuberculosis, realizing that most M. simiae and up to 2% or 3% of some other species may also give positive niacin reactions. Once the Niacin-Positive Nonchro- mogens are identified, one may refer to table 6A to determine how M. tuberculosis and M. simiae are identified. Colonies of M. tuberculosis always are rough, dry, and corded; on 7H-10 and other agar media, the colonies commonly are flat and spreading with an irregular edge, while on egg media the colonies often become more heaped in the centers (see color plate 23, page 147). This morphology, together with the use of niacin, nitrate reduction, and catalase activities enables the precise identification of 96% of M. tuberculosis (table 6B), while the 68°C catalase test and 131 ability to grow at 25°C will successfully identify 94% of M. simiae (table 7B). Besides the biochemical activities, M. simiae usually pro- duces smooth colonies after 2 to 3 weeks’ incubation on either egg- or agar-based media (color plate 24, page 147). Unless tests for niacin production and/or photochromogenicity suggest M. simiae, colonies of this taxon may be confused with M. avium complex. Strains of M. simiae often require prolonged (8 to 12 hours) light exposure and still longer (2 to 5 days) incubation to induce pigment production. Only rarely are M. tuberculosis and M. simiae confused with one another, and rarely are strains of other species confused with these two niacin- positive mycobacteria. TABLE 6A. Niacin-positive nonchromogens Test M. tuberculosis M. simiae* T° Range 33°-39° 22°-41° Nitrate Reduction + F Catalase, R® oo < 45mm > 45mm Catalase, 68°C — + Photoactive Pigment — + *Note, only niacin-positive M. simiae are included in this table (i.e., 85% of the strains of this species). The remaining niacin-negative isolates (15% of strains) are dealt with in tables 7A or 8A. TABLE 6B. Problems in differential identification—TB complex Subject Species, Possible Additional Key Test Pattern, and “Look-Alike” Differential (Probability of Correct ID) Species’ Tests? M. bovis 68° catalase _ M. tuberculosis Niacin/Nitrate reduction Growth on TCH — PZAase (0.87) M. tuberculosis Niacin production + Nitrate reduction + None 68° catalase en (0.96) 14 Look-alike” species arranged in order of decreasing likelihood of confusion with ‘‘subject species."’ If species name is followed by asterisk (*), then look-alike species may still be confused with subject species even after use of additional differential tests, 2The “additional tests’ are commonly arranged in order of decreasing differential value. Those enclosed in parentheses are (a) available only in a few reference laboratories, (b) not done routinely, or, (c) in a few cases, of limited value. 132 In fact, no species are likely to be confused with M. tuberculosis (table 6B) and only three species have the statistical probability to be con- fused with niacin-positive M. simiae (bottom table 7B). Pigment pro- duction will rule out most strains of M. gordonae that resemble M. simiae, while the semiquantitative catalase test helps with look-alike strains of M. avium. For both these other species, the use of seroag- glutination, available only from a select few reference laboratories, may be used for especially problematic cases. For the few strains of M. chelonae that give the same reaction pattern as niacin-positive M. simiae, growth rate and 3-day arylsulfatase will facilitate the separa- tion. Within the TB complex (table 6B), M. bovis is usually identified (87%) by a negative reaction in 68°C catalase and lack of growth on thiophen-2-carboxylic acid hydrazide (TCH). Colonies of M. bovis grown on 7H-10 or other oleic acid agar media are flat, rough, and corded much like M. tuberculosis; but when grown on egg media, the colonies commonly are thin, transparent, smooth, and pyramidal, with a slightly granular surface (see color plate 25, page 148). The only other species that is seen frequently enough in most laboratories to offer the statistical possibility of confusion with M. bovis is M. tuber- culosis. This has become even more of a problem in recent years with the influx of Asian immigrants. A number of recent isolates of M. tuberculosis from Cambodian refugees (and from Guamanians who may have been exposed to this same reservoir) have been shown to be inhibited by TCH, and thus to resemble M. bovis. Tests (repeated, if necessary) for niacin, nitrate reduction, and pyrazinamidase will obviate the problem. Also included in this group is the occasionally niacin-positive nonchromogen M. africanum. At present, this organism is seen pre- dominantly in central and western Africa as a cause of “tuberculosis” in humans, but the ease of world travel could alter this distribution in the future; already several isolates have been reported from western Europe and the United States. Properties of M. africanum are inter- mediate between M. tuberculosis and M. bovis. Growth is slow (often >40 days) and colonies are flat, rough, and dysgonic, but more eugonic growth is encouraged by the addition of 0.5% pyruvate to the isola- tion medium (2a). Mycobacterium africanum is commonly negative in tests for niacin production and nitrate reduction, but strains posi- tive in either or both these tests have been reported. Retention as a separate species is still in question, and further study is needed to establish the exact taxonomic position of M. africanum. B. Photochromogens Tests for the differentiation of the four potentially pathogenic Photochromogens are listed in table 7A, and the reliability of the diagnoses and the list of possible look-alike species are shown in table 7B. 133 TABLE 7A. Photochromogens Test M. kansasii |M. marinum| M. simiae | M. asiaticum Tween Hydrolysis + + — * Nitrate Reduction oo | CL — _ Catalase > 45mm os V + § Niacin _ eT Pyrazinamidase 4-day I. 3 — (some +) ? Arylsulfatase 2-week a + + . —/+ —/* Urease ¥ 1 . + _ In 88% of the cases, M. asiaticum may be identified by positive tests for photochromogenicity and Tween hydrolysis, coupled with nega- tive reactions for niacin and nitrate reduction (table 7B). Colonies are usually smooth and dysgonic after 2 to 3 weeks, rarely rough; they are usually photochromogenic, but occasional strains fail to turn yel- low after light exposure (color plate 26, page 148). The occasional strain of M. marinum that gives the same reactions as M. asiaticum may usually be identified first by the commonly superficial localiza- tion of M. marinum infections and second by different urease reac- tions. Although superficial localization of a disease process due to a slowly growing photochromogenic mycobacterium should always suggest M. marinum as the causative agent, this is not an irrefutable statement; on rare occasions, other photochromogens may be associ- ated with superficial disease in humans. Mycobacterium gordonae is the only other species that presents a possible source of confusion with M. asiaticum; this commonly is due to those rare strains of M. gordonae that may be pale yellow in the dark and exhibit pigment intensification after light exposure. If reexamination of pigment does not reveal a deeper yellow in the dark-grown M. gordonae, the laboratorian may have to search out a reference laboratory that per- forms serologic taxonomy tests. Mycobacterium kansasii is properly identified 96% of the time (table 7B) and, at least theoretically, no other species are seen in sufficient number to present a source of confusion for the laboratory worker. Colonies vary from flat, smooth to rough, raised, with irregular edges, as M. tuberculosis (see color plate 27, page 149). Although nonpig- mented when'grown in the dark, M. kansasii turns lemon yellow after exposure to light; prolonged light exposure (lighted incubator) may induce production of dark red crystals of B-carotene seen on the surface of the colonies. 134 TABLE 7B. Problems in differential identification—photochromogens Subject Species, Possible Additional Key Test Pattern, and ““Look-alike” Differential (Probability of Correct ID) Species’ Tests? M. asiaticum Tween hydrolysis + Nitrate reduction — Niacin production — Photochromogen + (0.88) M. kansasii Tween hydrolysis Nitrate reduction Niacin production - Photochromogen + (0.96) M. marinum Tween hydrolysis + M. Nitrate reduction — 2-wk Arylsulfatase + M. Photochromogen + (0.93) M. simiae (+ Photochromogen) M. Nitrate reduction Tween hydrolysis (Niacin production) + 0.72 (0.61, with niacin) M. simiae M. marinum Localization/Urease M. gordonae Recheck pigment/ (Serology) No species gordonae Localization/Urease asiaticum Localization/Urease avium Semiquant. catalase/ Urease (Serology) M. scrofulaceum (Nonphotochromogenic) Semiquant. catalase > 45 mm Iron uptake/Growth rate Growth rate/3-day Aryl/ 2-wk Aryl Growth @ 25° C/68°C Catalase M. fortuitum M. chelonae Tween hydrolysis M. tuberculosis (Niacin production) + M. avium Urease/ (Serology) 0.84 (0.72, with niacin) Other Rapid Growers | Growth rate M. simiae (Niacin +) M. gordonae Pigment/ (Serology) 68° C Catalase + M. avium Semiquant. catalase/ (Serology) Growth at 25° C + M. chelonae Growth rate/3-day Aryl (0.94) 14Look-alike" species arranged in order of decreasing likelihood of confusion with ‘subject species.” If species’ name is followed by asterisk (*), look-alike species may still be confused with subject species even after use of additional differential tests. 2The “additional tests’ are commonly arranged in order of decreasing differential value. Those enclosed in parentheses are (a) available only in a few reference laboratories, (b) not done routinely or (c) in a few cases, of limited value. 135 Mycobacterium marinum is identified (93% of the time) by a nega- tive nitrate reduction coupled with positive reactions in Tween hydrolysis, photochromogenicity, and 2-week arylsulfatase. Colonies may be smooth to rough on various egg media, but more commonly smooth on oleic acid agars (see color plate 28, page 150). On primary isolation (superficial lesions of humans) M. marinum exhibits more restricted growth temperature (25 to 33°C) but quickly adapts to growth at 37°C. As in the case of M. kansasii, the photochromogenic yellow pigment of M. marinum is easily seen on young, actively growing cultures exposed to light. Separation from occasional look-alike strains of M. asiaticum and M. gordonae may be accomplished, in both instances, by the urease test and determination of the location from which the organism was isolated (table 7B). Mycobacterium simiae is perhaps the most ill-defined species in the group of Photochromogens. In the bottom portion of table 7B, this species is presented in the three confusing forms in which it is most frequently seen. The niacin-positive strains are identified most reliably (94%), whereas the three look-alike species (M. gordonae, M. avium, and M. chelonae) are distinguished from M. simiae by pigment, semiquantitative catalase, and growth rate, respectively. The nonphotochromogenic M. simiae may be properly identified 84% of the time with additional tests for semiquantitative catalase and Tween hydrolysis. Nonetheless, look-alike strains are observed among M. tuberculosis, M. avium, and several rapid growers. Tests for growth rate, iron uptake, and arylsulfatase facilitate the separa- tion from the rapid growers. Mycobacterium tuberculosis may be ruled out by culture incubation at 25°C and the 68°C catalase test. If urease alone will not enable distinction from M. avium, one may have to resort to seroagglutination studies. The photochromogenic M. simiae are most difficult to identify (72%) exactly; the look-alike M. avium generally may be ruled out with tests for catalase and urease. If more exact pigment production does not enable the separation from M. scrofulaceum, seroagglutination may have to be utilized. See Niacin-Positive Nonchromogens for com- ments on pigment production. The most striking thing about M. simiae is the difficulty of precise identification (table 7B). This is further reinforced by the fact that some strains of M. asiaticum were among the early “cluster” of organ- isms named M. simiae. Although most clinically important species of Mycobacterium exhibit probabilities of correct identification in excess of 90%, the pigment variants of M. simiae show only 72% and 84% likelihoods for correct identification. This may be due to extreme test variability for the species; but our past experience suggests that an equally plausible explanation is the possibility that the species’ may still be a mixture of two or more separate species or subspecies. Only time will resolve this problem. 136 C. Nonphotochromogens An examination of figure 19 shows that the Tween hydrolysis test provides a fairly reliable method of separating potentially pathogenic (negative) from commonly saprophytic (positive) nonphotochromo- genic mycobacteria. In fact, the test would have had even greater value in the 1975 edition of this manual (10) before the potential pathogen M. malmoense was recognized. Figure 19 shows the reli- ability of the Tween hydrolysis test when examining two different sets of culture data. One set represents a very select population of 864 strains of mycobacteria submitted to CDC (figure 19A) for confir- mation of identification. In this array of results, only 85% of poten- tially pathogenic mycobacteria gave negative reactions in Tween hydrolysis, whereas 15% behaved like saprophytes and yielded posi- tive reactions. Almost all the discrepant results were due to positive reactions by a disproportionately high number of cultures of M. malmoense. Most submissions of this relatively new Mycobacterium to the CDC reflect the uneasiness of other reference laboratories in making a specific identification of the new species. On the other hand, when reliability of Tween hydrolysis is examined by using the 41,291 cultures examined by the State health departments, it is noted (figure 19B) that test reliability climbs to 99% for both potential pathogens and common saprophytes. This is due almost totally to the fact that only about 20 strains of M. malmoense were reported nationally. This represents <0.05% of the potentially pathogenic nonphotochromogens (table 3), and their few positive reactions in Tween hydrolysis are negligible in the face of nearly 7000 M. avium complex, most of which yielded negative reactions. When used on a national basis then, the Tween hydrolysis test is still highly reliable for the separa- tion of potential pathogens from saprophytes among the nonphoto- chromogens. A. CDC Data B. State Health Department Data Percent Reacting as Percent Reacting as Actual Actual Classification | Pathogens |Saprophytes Classification | Pathogens |Saprophytes Pathogens 85* 15 Pathogens 99* 1 Saprophytes 2 98 Saprophytes 1 99 * Low positive due to Tweent+ M. malmoense *Test improved because few M. malmoense seen FIGURE 19. Nonphotochromogens—separation of pathogens and saprophytes using Tween Hydrolysis Test 137 Tests for the separation of clinically important Nonphotochromo- gens are given in table 8A. Table 8B records both the reliability of the specific identifications and the list of look-alike species that may be a source of confusion for differential identification. Mycobacterium avium (this includes M. intracellulare) is relatively nonreactive in most of the differential biochemical tests, with 94% of strains producing <45 mm of foam in the semiquantitative catalase test and negative reactions in tests for Tween hydrolysis, X-colony form, niacin production, and 3-day arylsulfatase (table 8B). Colonies of M. avium reveal extreme variability. From diseased humans, the initial growth is often smooth, thin, very transparent, and difficult to see. On occasion, rough colonies similar to M. tuberculosis are seen. On repeated passage, these colonies tend to become smooth, hemi- spherical, and opaque (see color plate 29, page 151). Look-alike spe- cies include M. tuberculosis, M. scrofulaceum, M. bovis and “other” rapid growers. The latter may be differentiated by growth rate and, if necessary, urease activity. Mycobacterium tuberculosis is distin- guished (even if niacin-negative) by positive test reactions for nitrate reduction and urease. Mycobacterium scrofulaceum is commonly seg- regated by pigment production, but when that feature is not definitive, reaction in the urease test usually is. If both these last two features fail, serologic agglutination tests may be needed. The very nonreac- tive M. bovis often presents differential problems, and it is only its TABLE 8A. Nonphotochromogens o |& x| § i © § = « $5885 83 & |.§.% Q Se < © Sa £ 3 a Q | 3 £ 2 o> |g 5 Sf Ss 3 2 So|s S |s38 s g 3 Test < Catalase, R° <45 | <45 |[<45 | <45 | >45 >45 <45 — Catalase, 68°C —_ + + _— + + Vv _ Tween Hydrolysis — — ee + + — + _ Tellurite Reduction | — + — ir ro wwf: + — Nitrate Reduction —_ — mm — + — — _ Growth on TCH — + + + + + + | not done NaCl Tolerance _ — — — |—/+ ft _ _ Urease + — — + — + _ _ Pyrazinamidase 4-day/7-day —_——| + + |[—/F |F/+ Vv “+ + Photoactive (not intense) Pigment — _—_ —_— — — + _ _ Niacin —_— — _— — mn + (some—) a — See Table 9A for description of MAIS intermediates. 2 Grows 22-37 °C, one serotype, usually smooth, domed colonies, susceptible EMB, ETA, KM, CS. 3 Grows 25-30 °C, not at 37°C, requires hemin for growth; colonies usually rough. 138 paucity in numbers isolated that prevent it from being a more perplex- ing organism when it comes to separation from M. avium. Tests for susceptibility to TCH and urease activity commonly enable the defini- tive speciation of these two organisms. TABLE 8B. Problems in differential identification—nonphotochromo- gens Subject Species, Possible Key Test Pattern, and ‘“’Look-alike’” Additional Differential Tests? (Probability of Corerct ID) Species! M. avium Tween hydrolysis - M. tuberculosis Urease/Nitrate reduction Semiquant. catalase < 45mm |M. scrofulaceum Pigment/Urease/(Serology) Niacin production — M. bovis | TCH/Urease 3-day Arylsulfatase — Other Rapid Growers Growth rate/Urease ““X"" colony3 - | (0.94) | M. gastri Semiquant. catalase <45 mm Nitrate reduction — M. avium Urease/68° C catalase “X" colony — Other Rapid Growers Growth rate Tween hydrolysis + 2-wk Arylsulfatase + Nonphotochromogen + (0.94) M. malmoense M. gordonae™ Pigment/ (Serology) Semiquant. catalase < 45 mm | M. gastri 2-wk Aryl/PZAase (4-day)/ (Serology) Tween hydrolysis + | M. tuberculosis Growth @ 25° C/Niacin Nitrate reduction _ M. avium (Morphology/Serology) “"X" colony —- M. marinum Pigment (0.94) Other Rapid Growers Growth rate M. terrae cplx | M. gordonae* Pigment/ (Serology) Tween hydrolysis + M. kansasii Pigment/Urease/ (Serology) Semiquant. catalase > 45mm | M. tuberculosis Niacin/68° C catalase “'X" colony —- M. asiaticum Pigment/Nitrate reduction Sodium chloride — M. flavescens Pigment/Tween opacity 3-day Arylsulfatase - Other Rapid Growers Growth rate /Urease (0.90) M. szulgai Pigment/Urease M. triviale Semiquant. catalase > 45 mm | M. fortuitum Growth rate/MacConkey Tween hydrolysis + M. chelonae (abscessus) | Growth rate/Nitrate reduction Sodium chloride + Other Rapid Growers Growth rate/Tellurite reduction Nonphotpchromogen + M. terrae cplx 3-day Aryl/ (Morphology) (0.90) 1 “| ook-alike’ species arranged in order of decreasing likelihood of confusion with “subject species.” If species’ name is followed by asterisk (*), look-alike species may still be confused with subject species even after use of additional differential tests. 2 The “additional tests’ are commonly arranged in order of decreasing differential value. Those enclosed in parentheses are (a) available only in a few reference labora- tories, (b) not done routinely or, (c) in a few cases, of limited value. 3 ux" refers to the compact, smooth-to-rough, slow-growing colonies that often ex- hibit tiny stick-like projections at the periphery when examined under transmitted light. Their uniqueness to M. xenopi is responsible for the designation ““X". 139 The commonly saprophytic M. gastri is identified 94% of the time by positive tests for Tween hydrolysis, 2-week arylsulfatase, and lack of pigment, coupled with negative reactions for nitrate reduction, X-colony formation, and production of less than 45 mm of foam in the catalase test (table 8B). Colonies on egg media are smooth to rough, nonpigmented, whereas those on oleic acid-agars more com- monly are smooth, domed, often resembling the smooth colony type of M. kansasii (color plate 30, page 152). The look-alike M. avium may be reliably separated from M. gastri by urease and 68°C catalase tests, whereas the simple observation of rate of growth is adequate to separate M. gastri from the “other rapid growers.” The relatively new M. malmoense may be correctly identified 94% of the time by positive Tween hydrolysis, less than 45 mm of foam in the catalase test, and negative observations in X-colony formation and nitrate reduction. Colonies are usually smooth (color plate 31, page 152). Because of the large numbers of isolates seen, neither M. gordonae nor M. avium may be separated from M. malmoense with absolute certainty in spite of additional features of pigment and morphology; serology may be the only truly helpful test. Among the other look-alikes of M. malmoense, M. gastri is separated by 2-week arylsulfatase and pyrazinamidase; M. tuberculosis, by growth rate at 25°C and niacin test; M. marinum, by pigment; and other rapid growers, by growth rate (table 8B). The M. terrae complex also contains M. nonchromogenicum (and occasionally M. triviale, if growth on NaCl is disregarded); but a posi- tive Tween hydrolysis, more than 45 mm of foam in the catalase test, and negative reactions in X-colony formation, sodium chloride tolerance, and 3-day arylsulfatase will properly identify 90% of the complex (exclusive of M. triviale). Colonies vary from thin, flat, and smooth to rough (color plate 32, page 153); they are usually nonpig- mented, but pale pastel colors (beige, yellow, pink) are occasionally seen. Mycobacterium gordonae cannot always be distinguished from M. terrae even with pigment formation, so serology may be needed to effect the precise separation. Pigment and urease will enable the distinction of M. terrae from M. kansasii and M. szulgai. M. tuberculo- sis is separated by niacin and 68°C catalase; M. asiaticum by pigment and nitrate reduction. Mycobacterium flavescens is differentiated by pigment production and Tween opacity; the “other” rapid growers are separated by growth rate and urease. If grown at 37°C rather than 28°C, some M. chelonae may react as M. terrae, so incubation tempera- ture is quite important. Mycobacterium triviale is a very reactive nonphotochromogen, with 90% of isolates expressing nonpigmented (rough corded) colonies with hyperactive (>45 mm) catalase, positive reactions in Tween hydrolysis and tolerance of sodium chloride (NaCl). Members of the M. fortuitum complex and “other” rapid growers may be separated 140 from M. triviale by combinations of growth rate, MacConkey agar, nitrate reduction, and tellurite reduction. The closely similar M. terrae may generally be separated by 3-day arylsulfatase and colony mor- phology (table 8B). Other nonphotochromogenic slow growers listed in table 4 or referred to only casually in the text need some discussion. In extended taxonomic studies, M. intracellulare and M. avium may be segre- gated into separate but overlapping clusters (11); however, no bio- chemical tests are sufficiently definitive to facilitate their precise separation by the clinical microbiology laboratory, so the two species commonly are lumped together as M. avium complex. Occasional strains of mycobacteria that exhibit variable reaction patterns in tests for pigment, urease, Tween hydrolysis, and catalase that are inconsis- tent with either M. avium, M. intracellulare, or M. scrofulaceum, have been lumped into the so-called MAIS intermediates (see also table 9A for variations most often seen at CDC). The MAIS group contains only strains truly intermediate in properties of the three species from which the acronym derives; strains that may be specifically identified as one or another of the three species above should not be categorized as MAIS intermediates. Mycobacterium shimoidei is considered to be pathogenic for humans (11), but little is known of its biochemical variability because so few strains have been seen. Examination of table 4 reveals the few known properties of M. shimoidei to be very similar to those of both M. avium and M. malmoense. A closer examination of more strains TABLE 9A. Scotochromogens 3 . [) 2 8 g 3 g g . 3 > Q S a n 3 5 £ © o = = N Q ~ > << 2 3 x g 3 s Q x s > ~ 12] x s = = Test Tween Hydrolysis — |+ (some slow)| — + + _— + Nitrate Reduction —_— + — — +/— | — — Catalase, R° >45 >45 <45 >45 >45 |<45 | <45 NaCl Tolerance —_— _ — _ + —_ —_ Urease +(—) + — | — (some +)| + + — Photoactive Pigment 25°C — # — — it = pas 2-Week Arylsulfatase | —/+ —/wk + + —/ + + x — Tween Opacity so —1w/+5w — —_ +/1w | — — * MAIS or MAIS intermediates exhibit variable properties that are a combination of those seen in M. avium, M. intracellulare, M. scrofulaceum. The properties listed for MAIS intermediates are those most commonly observed at CDC; other combinations of test reactions may be seen. 141 will be needed to determine if this taxon is either (a) truly distinct or (b) seen with sufficient frequency to be a source of taxonomic confu- sion to the clinical microbiologist. Colonies are commonly rough, non- pigmented after 2- to 3-weeks’ growth. Mycobacterium ulcerans causes ulcerative cutaneous or subcutane- ous lesions in humans and has never been isolated from sputum. The organisms seem to be confined to tropical or subtropical climates but, on rare occasions, may be encountered in travelers from such areas (e.g., diplomatic persons, Peace Corps workers). The ulcerous lesion is characterized by a deeply undermined edge that increases in size by direct extension to involve large areas around the initial site of infection. Mycobacterium ulcerans grows extremely slowly (6 to 9 weeks) at a sharply restricted temperature range (30 to 33°C). Colonies, tiny and transparent at first, become low, flat, rough, and range from nonpigmented to very pale yellow. Other properties are as described in table 4. Mycobacterium haemophilum causes skin lesions in humans, par- ticularly those undergoing immunosuppressive therapy. The species has a rather narrow optimum growth range (30 to 33°C) and yields nonpigmented rough (rarely smooth) colonies after 3 or 4 weeks’ incubation on either egg- or agar-based media supplemented with 0.4% hemoglobin or 60 pM hemin (11). Because 15 pg/ml ferric ammonium citrate may be substituted for hemin, the medium used for the iron uptake test may be used for primary isolation of this organism; both blood agar and chocolate agar have also been used for primary isolation. The last nonphotochromogen that needs mention is the usually saprophytic M. nonchromogenicum, commonly included in the M. terrae complex. Tests that offer some consistency for separation of M. nonchromogenicum from M. terrae are those for pyrazinamidase and nitrate reductase. Mycobacterium nonchromogenicum is com- monly positive in PZAase and negative in nitrate reductase. Because all members of the M. terrae complex are usually saprophytic, most clinical laboratories tend not to attempt separation beyond the com- plex level unless the isolate is one of those that only rarely is associ- ated with disease. D. Scotochromogens Just as for the Nonphotochromogens, the Tween hydrolysis test was early reported to offer a preliminary separation of potential patho- gens from saprophytes among the Scotochromogens. Figure 20 again shows the value of the test is increased depending upon the source of data examined. When the unusual assemblage of strains submitted to CDC was examined, reliability of the Tween hydrolysis test for identification of potential pathogens was only 84%, the main reason 142 A. CDC Data B. State Health Department Data Percent Reacting as Percent Reacting as Actual Actual Classification | Pathogens |Saprophytes Classification | Pathogens |Saprophytes Pathogens 84 16% Pathogens 95 b5* Saprophytes 1 99 Saprophytes 1 99 *M. szulgai often Tween hydrolysis positive. Because few strains seen nationally, reliability of test is increased. FIGURE 20. Scotochromogens—separation of pathogens and sapro- phytes using Tween Hydrolysis Test for poor predictability being the disproportionate number of M. szulgai, often positive in Tween hydrolysis. In contrast, when national data from the State health departments (SHD) were assessed, test reliabil- ity for prediction of pathogenicity was raised to 95%, primarily because of the very low numbers of M. szulgai seen nationally. Once again, as awareness of new species is increased, reliability of the Tween hydrolysis test may decrease. Whether or not the Tween hydrolysis test is used for preliminary subdivision of Scotochromogens, the definitive identification may be accomplished as shown in table 9A. The reliability of the species identification and the problem “look-alike” organisms are listed in table 9B. Separation within the Scotochromogens has been especially diffi- cult at CDC mainly because of the strange and unpredictable reac- tions seen in the test for urease. So unreliable have been the results that the test is not noted as “’key’’ in any of the definitive properties of table 9B, but it is listed as an ancillary test, whereas the data in table 4 reflect the urease reactions commonly reported by others for this test. Reasons for the failure of CDC to obtain reliable test patterns in urease reactivity are not known but may include (a) the highly selec- tive population of organisms sent to CDC, (b) the test method(s) used here, or (c) some error in test performance. Because of this failure, the nitrate reduction, Tween hydrolysis, catalase, and Tween opacity tests are the ones that afford separation of the species. Mycobacterium flavescens may be grouped either with Rapid Grow- ers or with Scotochromogens, depending on the speed of growth. The only organisms that are seen in sufficient number to cause great confusion are the “other” Rapid Growers. Rapid growth (<5 days) will usually separate the true Rapid Growers, whereas well-diluted MV. flavescens commonly takes > 7 days to yield growth of well-isolated, 143 mature colonies. If M. flavescens is properly listed as a slow grower, it is properly identified 90% of the time (table 9B); if over inoculation results in more rapid growth, definitive identification is more difficult. Colonies commonly are soft, smooth, yellow-orange, and butyrous after more than 7 days. TABLE 9B. Problems in differential identification—scotochromogens Subject Species, Key Test Pattern, Possible iti ifferenti 2 and (Probability of Correct ID) ““Look-alike” Species! Additional Differential Tests M. flavescens Tween hydrolysis Nitrate reduction Tween opacity “X"" colony3 Scotochromogen (0.90) M. gordonae Tween hydrolysis + Semiquant. catalase > 45 mm [Other Rapid Growers [Growth rate Nitrate reduction —- “X"" colony — Tween opacity — Scotochromogen + (0.84) M. scrofulaceum Tween hydrolysis — M. gordonae Urease/ (Serology) 3-day Arylsulfatase — ““X"" colony — Other Rapid Growers | Growth rate/MacConkey agar Semiquant. catalase > 45mm Scotochromogen + (0.84) M. szulgai Nitrate reduction + Semiquant. catalase/ > 45 mm | M. gordonae * Urease/T*-dependent photochrome 3-day Arylsulfatase - “X" colony - M. scrofulaceum T "dependent photochrome/ (Serology) Tween opacity - Scotochrome 35°C + Other Rapid Growers | Growth rate (0.94) M. xenopi Semiquant. catalase < 45 mm Tween hydrolysis - M. avium 3-day Aryl/ (Serology) Niacin production — Photochromogen - “X" colony + (0.95) Other Rapid Growers" [Growth rate + + + ' “Look-alike’’ species arranged in order of decreasing likelihood of confusion with subject species. If species name is followed by asterisk (¥), look-alike species may still be confused with subject species even after use of additional differential tests. 2 The ‘additional tests’’ are commonly arranged in order of decreasing differential value. Those enclosed in parentheses are (a) available only in a few reference laboratories, (b) not done routinely or, (c) in a few cases, of limited value. 3X" refers to the compact, smooth-to-rough, slow-growing colonies that often exhibit tiny stick-like projections at the periphery when examined under transmitted light. Their uniqueness to M. xenopi is responsible for the designation ““X". 144 Mycobacterium gordonae is probably the most common sapro- phytic Mycobacterium recovered in the mycobacteriology laboratory. It is found in a variety of environmental sources and quite commonly in water taps where it may ultimately generate problems of “‘false- positive’ smears, unexpected positive cultures, and occasionally mini- "pseudoepidemics’’ in laboratories whose water supply may have been excessively contaminated with M. gordonae because of heavy rains and resultant overtaxed water purification systems. It is recog- nized (84%) by >45-mm semiquantitative catalase and a positive Tween hydrolysis, whereas a negative Tween opacity test helps to separate it from M. flavescens. Colonies are usually smooth, yellow to orange; pigment may intensify if cultures are exposed to light continuously (color plate 33, page 154). In those laboratories where the urease test works, a negative reaction for M. gordonae helps in its separation from M. scrofulaceum. It should be remembered, however, that urease-positive M. gordonae are known (var. ureolyticum). Possible confusion with Rapid Growers other than M. fortuitum complex is obviated by a check on rate of growth. As shown in table 9B, M. scrofulaceum is identified (84%) by its general lack of reactivity in all the tests listed and its production of >45-mm of foam in the semiquantitative catalase test. It is most com- monly found in pus from draining cervical lymph nodes of children; much more rarely in sputum. Culture growth on both egg- and agar- based media is usually yellow to orange, smooth, pyramidal, more rarely rough; on subculture, colonies tend to become more domed (color plate 34, page 154). The fact that most other laboratories obtain positive urease tests for M. scrofulaceum should be explored as yet another differential feature that, if reflected in the broader range of M. scrofulaceum seen "in the field,” may be invaluable in separating this potential pathogen from the much more common saprophyte, M. gordonae. Mycobacterium szulgai, another of the more recent potential patho- gens and still rather uncommon, is usually recognized (94%) by its very strong nitrate reductase activity. A less consistent feature is the ability of M. szulgai to express a photochromogenic pigment when grown at 25°C, while being scotochromogenic at 35 to 37°C. This special photoactive pigment and a ““good’’ urease test help to sepa- rate M. szulgai from M. scrofulaceum and M. gordonae, respectively. On growth medium, either smooth or rough colonies may be seen (color plate 35, page 155). Mycobacterium xenopi is found on the charts of both Nonphoto- chromogens (table 8A) and Scotochromogens (table 9A), and its unique characteristics (except for the lack of a singular kind of pigment) make it quite easy to identify (95%), regardless of which pigment- group it falls into. Among the Scotochromogens, the almost unique features are very slow growth (>4 weeks), less than 45 mm of foam in 145 the semiquantitative catalase test, together with positive results in 3-day arylsulfatase (liquid substrate) and presence of the “X" colony on agar medium (e.g., 7H-10). If the 3-day arylsulfatase test does not aid in the separation of M. xenopi from M. avium look-alikes, the laboratory may have to request serologic testing as a differential aid. See color plate 35D, page 155. Although it is not possible to identify precisely all acid-fast bacilli, a reasonably exact identification may be attained by (a) careful atten- tion to the figures and tables in this chapter, (b) knowledge of the colonial morphologies of the organisms on both egg- and agar-based media, and (c) judicious use of the predictability charts (both for species identification and frequency of species isolation). Expertise in differentiation of mycobacteria is attained by experience with large numbers of strains. With this experience comes a sagacity, or “gut feeling,” about these organisms that cannot possibly be incorporated into the predictability charts but, nevertheless, ultimately turns out to be a major contribution to the final identification of an unknown Mycobacterium. REFERENCES 1. Good RC. Isolation of nontuberculous mycobacteria in the United States, 1979. J Inf Dis 1980;142:779—83. 2. Good RC, Snider DE Jr. Isolation of nontuberculous mycobacteria in the United States, 1980. J Inf Dis 1982;146:829 33. 2a. Huet M, Rist N. Advantage of using pyruvate media for the culture of tubercle bacilli in Black Africa. Bull Int Un Tuberc 1973;48:91-5. 3. Kubica GP, Gross WM, Hawkins JE, Sommers HM, Vestal AL, Wayne LG. Laboratory services for mycobaterial diseases. Am Rev Respir Dis 1975;112:773—-87. 4. Runyon EH. Mycobacteria encountered in clinical laboratories. Leprosy Briefs 1958;9:21-3. 5. Runyon EH. Anonymous mycobacteria in pulmonary disease. Med Clin North Am 1959;43:273-90. 6. Runyon EH. Pathogenic mycobacteria. Adv Tuberc Res 1965;14:235—-87. 7. Runyon EH. Ten mycobacterial pathogens. Tubercle 1974;55:235—40. 8. Runyon EH, Karlson AG, Kubica GP, Wayne LG . Mycobacterium. In: Lennette EH, Spaulding EH, Truant JP. eds. Manual of clinical microbiology, 2nd ed. Washington, D.C: American Society for Microbiology. 1974:148—-74. 9. Timpe A, Runyon EH. The relationship of ““atypical’’ acid-fast bacteria to human disease. J Labor Clin Med 1954;44:202—-9. 10. Vestal AL. Procedures for the isolation and identification of mycobacteria. Atlanta: Centers for Disease Control, PHS, HEW, 1975. 11. Wayne LG, Kubica GP. Family Mycobacteriacease In: Holt JG, Sneath PHA, eds. Bergey's manual of determinative bacteriology, 9th ed. Baltimore: Williams and Wilkins, 1985, in press. 146 Cc COLOR PLATE 23. M. tuberculosis. a. Rough colony on 7H-10 agar. b. Same colony on 7H-10 agar photographed with transmitted light to show cords. c. Rough, dry, heaped up in the center colony on egg medium. A COLOR PLATE 24. M. simiae. a. Smooth dome colonies on 7H-10 agar grown in the dark, photographed with a blue filter. b. Smooth dome colonies exposed to light, grown on egg medium. 147 Cc COLOR PLATE 25. M. bovis. a. Rough colonies on 7H-10 agar photographed with green filter. b. Rough, corded colonies on 7H-10 agar photographed with transmitted light. c. Rough, dry colonies on egg medium. d. Smooth pyramid colonies on egg medium. A COLOR PLATE 26. M. asiaticum. a. Smooth dome colonies exposed to light grown on 7H-10 agar, photographed with blue filter. b. Smooth dome colonies exposed to light grown on egg medium. 148 COLOR PLATE 27. M. kansasii. a. Rough, dry colonies with irregular edges, grown on 7H-10 agar photographed with blue filter. b. Rough colonies on 7H-10 agar photo- graphed with transmitted light. c. Rough nonpigmented colonies on egg medium grown in the dark. d. Smooth dome yellow colonies with thin, irregular edge on 7H-10 exposed to light and photographed with a green filter. 149 COLOR PLATE 28. M. marinum. a. Irregular pigmented colonies on 7H-10 agar exposed to light photographed with a blue filter. b. Irregular pale yellow pigmented colony with a thin, trailing edge, grown on egg medium and exposed to light. c. Irregular lobular colonies grown in the dark on egg medium. 150 COLOR PLATE 29. M. avium complex. a. Rough colonies on 7H-10 photographed with green filter. b. “TB-like” colonies on 7H-10 photographed with transmitted light. c. Smooth dome large and small colonies on egg medium. d. Small and large colonies on 7H-10 agar photographed with a blue filter and transmitted light. e. Several colony forms on 7H-10 photographed with transmitted light. f. Smooth dome colonies, thin transparent colonies and central rough colony with lobular dome. This mixed colony morphology is typical for M. intracellulare. 151 COLOR PLATE 30. M. gastri. a. Smooth dome nonpigmented, small and large colo- nies on egg medium. COLOR PLATE 31. M. malmoense. a. Three smooth colonies with central dome and one irregular rough colony also with central dome on 7H-10 photographed with blue filter. b. Same four colonies on 7H-10 agar photographed with transmitted light to enable better detection of the different colony forms. 152 ee oo o® eo, E COLOR PLATE 32. M. terrae complex. a. M. terrae — smooth dome colonies with thin trailing edge on egg medium. b. M. terrae colonies on 7H-10 agar photographed with transmitted light and a green filter to better demonstrate the thin, trailing edge. c. Rough, dry colonies of M. triviale on egg medium. d. M. triviale on 7H-10 agar photographed with a blue filter. e. M. triviale on 7H-10 agar photographed with transmitted light. 153 B COLOR PLATE 33. M. gordonae. a. Pale yellow smooth dome colonies on 7H-10 agar photographed with blue filter. b. Orange smooth dome colonies on egg medium. B COLOR PLATE 34. M. scrofulaceum. a. Smooth dome yellow colonies on 7H-10 agar photographed with blue filter. b. Smooth dome yellow colonies on egg medium. 154 D COLOR PLATE 35. M. szulgai. a. Smooth dome and rough colonies on 7H-10 agar photographed with blue filter. b. Smooth dome and rough colonies on 7H-10 agar photographed with transmitted light. c. Smooth dome colonies on egg medium. d. M. xenopi colonies on 7H-10 agar. 155 COLOR PLATE 36. M. fortuitum. a. Donut colonies on egg medium. b. Rough, dry colonies on 7H-10 agar photographed with blue filter. c. Rough, dry colonies on 7H-10 agar photographed with transmitted light. d. Smooth colonies on 7H-10 agar with green filter. 156 COLOR PLATE 37. M. chelonae. a. Two smooth dome colonies with narrow, thin irreg- ular edges and two pyrimidal colonies with lobular centers on 7H-10 agar photo- graphed with green filter. b. Same four colonies on 7H-10 agar photographed with transmitted light. c. Smooth dome colonies on egg medium. 157 Antituberculosis Chemotherapy and Drug Susceptibility Testing Antituberculosis Chemotherapy and Drug Susceptibility Testing The early years of antituberculosis chemotherapy were plagued with treatment failure and relapse. The high risk of failure during the early years of drug treatment was due to the selection of drug-resistant mutants during the initial phase of single drug therapy (9,34), whereas the high rate of relapse after treatment was a result of the reemer- gence of viable organisms (both resistant and susceptible forms) that had persisted during drug treatment (32). Today, chemotherapy is aimed not only at prevention of both treatment failure and relapse, but also at complete sterilization of the lesion in the shortest time possible (16). To achieve this goal is the rationale for short course chemotherapy with multiple drug combinations. A. Preventive Therapy (or Chemoprophylaxis) for Tuberculosis Infection Preventive therapy is a form of treatment to minimize the chance for developing clinical disease after infection has occurred (19). Most people (ca. 90%) infected with tubercle bacilli remain well and will never develop clinical disease; but some (ca. 10%), without the bene- fit of preventive treatment, will eventually develop active tuberculo- sis and become the sources of new infections (12). If preventive therapy can be administered and completed, the risk of developing disease, among the 10% who will acquire tuberculosis, is reduced by 55% to 90%. Thus, preventive therapy not only benefits the health of the treated person but, in terms of public health, also minimizes the potential spread of tuberculosis. Preventive therapy for tuberculosis became a practical possibility in 1955 after the introduction of the safe, inexpensive, oral drug, isoniazid (INH) (13). Before preventive treatment is administered, cer- tain individualized factors should be considered: What is the estimated risk for developing clinical disease? What is the risk of developing a serious adverse reaction to isoniazid? What are the chances of infecting others? 159 Isoniazid preventive therapy is recommended for people consid- ered at high risk. Among those at high risk are: 1. Close contacts of a patient who has newly discovered tuberculosis 2. Tuberculin skin test reactors with evidence of dormant tuberculo- sis by roentgenogram 3. Newly infected persons; i.e., recent tuberculin converters 4. Infected persons with certain special clinical conditions. Some conditions that increase the risk of developing tuberculosis are: prolonged adrenocorticoid therapy, immunosuppressive therapy, some hematologic and reticuloendothelial diseases such as leu- kemia or Hodgkin's disease, silicosis, postgastrectomy, and bypass surgery for obesity (33) 5. Persons with significant tuberculin reactions, who are less than 35 years of age Preventive therapy with isoniazid is usually inadvisable for people over 35 years of age (because of the possiblity of age-related liver disease), unless they are in one of the four high-risk groups listed above. Isoniazid is also contraindicated among those who have a history of previous isoniazid-associated hepatic injury or allergic reactions, or who have acute liver disease of any cause. The currently recommended duration of preventive therapy is 12 months, but shorter treatment periods (e.g., 6 months) have a signifi- cant protective effect. The dosage is 300 mg of isoniazid per day for adults and 10 mg/kg (not to exceed 300 mg per day) for children. Before starting preventive treatment a chest film should be taken to rule out the possibility of clinically progressive disease. B. Chemotherapy for Disease The effectiveness of chemotherapy for tuberculosis is dependent upon many factors; among them: the tubercle bacillus and its meta- bolic activity, the size of the bacillary population in the lesion, the pharmacodynamics of the antituberculosis drugs, the site of the disease, and the compliance of the patient. The bacterial populations within the lesions of human tuberculosis vary considerably in the speed with which they grow and metabolize, depending upon the particular environment of the bacilli. The bacilli located in the cavity wall are actively multiplying because of the abundance of oxygen, whereas those located in closed lesions deprived of oxygen are likely to be more slowly metabolizing. Figure 21 presents the hypothesis of how different drugs act on the special populations of tubercle bacilli in the lesions (31). 160 Population A consists of those organisms that are actively multiply- ing. Isoniazid acts quickly to kill these organisms, whereas rifampin (RMP) and streptomycin (SM) show slightly lesser degrees of effec- tiveness. Isoniazid kills the most actively multiplying organisms, but as the rate of bacterial growth slows, isoniazid begins to lose its effectiveness. In human tuberculosis, Population A is the largest and is responsible for (a) the positive smears and cultures seen in the laboratory and (b) most of the drug-resistant mutants that arise dur- ing or after therapy. Population B, figure 21, consists of many slowly growing organisms found inside macrophages where the pH is acid (pH 5.5). Pyrazinamide (PZA) is most effective against these more slowly growing organisms in an acid environment. Population C rep- resents those dormant organisms that occasionally have short peri- ods (spurts) of active growth. Rifampin is remarkably effective against these organisms because of its ability to kill rapidly (within 15 to 20 minutes). Finally, Population D represents those organisms that are completely dormant. These organisms are unlikely to be killed by any drug. When a patient's disease relapses, even after effective treatment, it is primarily Population D and, to a lesser extent, Population C that is responsible. ACTIVE A ~— INH (RMP, SM) CONTINUOUS GROWTH (Extra-Cellular) PZA _ B Cc METABOLISM | 5cip ENVIRONMENT SPURTS OF METABOLISM D PERSISTORS (No Drugs) SLOW (Populations B, C, D are never seen in microscopy or culture.) (Adapted from Mitchison, ref. 31) FIGURE 21. Drugs affecting special TB populations 161 Some of the more effective short-course regimens of treatment exploit the unique antibacterial activities of drugs against the differ- ent microbial populations outlined in figure 21. An extremely effective 6-month regimen used in controlled trials in some developing nations of the world, and also recommended by the British Thoracic Association and the American College of Chest Physicians, is one in which INH, RMP, PZA, and either SM or ethambu- tol (EMB) are given, under supervision, daily for 2 months; for the final 4 months of treatment, only INH and RMP are given either daily or two to three times per week. Cures in controlled trials approach 100%. Relapses, if they occur, are commonly with organisms that are totally susceptible to the four drugs; this is because the relapse is due almost exclusively to the dormant Population D, a group of organ- isms that are not metabolizing and hence cannot be affected by the drugs that would select the resistant mutants. In any TB control program, bacteriologic monitoring of patient response to therapy is very important. Treatment regimens may need to be changed or prolonged if the patient's culture is not negative after the fourth or fifth month of therapy. C. Drug Susceptibility Tests 1. Difficult to Standardize Drug susceptibility testing is one of the most difficult procedures to standardize for the mycobacteriology laboratory. Proficiency in per- forming susceptibility tests demands an understanding of: (1) The origin of drug resistance (2) The variation in stability of drugs subjected to different conditions of filtration, heat, or freezer storage (3) The alteration in the antimycobacterial activity of certain drugs when incorporated into different kinds of media (4) The type of susceptibility test performed (5) The reading and reporting of test results (6) The criteria of resistance These points are thoroughly discussed in several publications (3, 4, 6, 7, 22) that the interested reader is urged to consult for greater detail. In the ensuing pages, each subject will be addressed primarily from the point of view of performance at CDC. 2. Who Performs Drug Susceptibility Tests? In the United States, the ‘Levels of Service’ concept introduced by Kubica & Dye (23) and supported by the American Thoracic Society (18) recommends that three levels of laboratory work performance be 162 established, based on work-load, expertise, cost-effectiveness, and interest. Drug susceptibility tests should be performed only in labora- tories that are proficient in identifying the species of Mycobacterium being tested. In accordance with the “Levels of Service” concept, the Level Il laboratories may or may not choose to perform drug suscepti- bility studies on the strains of M. tuberculosis isolated. Most small Level Il laboratories have infrequent opportunity to perform such tests; therefore, proficiency and cost-effectiveness may best be main- tained in those large Level ll or Level lll laboratories where at least 10 drug susceptibility test patterns are done each week. 3. When to Perform Drug Susceptibility Tests The American Thoracic Society (1a) sets forth two options in deter- mining the need for susceptibility tests on initial isolates from a patient. The first option is to test all initial isolates. The second is to test isolates of (a) patients in high-risk groups for primary drug resistance (i.e., immigrants from certain high prevalence areas, contacts of known resistant cases, and residents of geographic areas where high levels of drug resistance have been documented) and (b) patients with life- threatening illness such as widely disseminated disease or meningitis. Generally, in communities where the incidence of primary drug resis- tance is less than 5%, there is no need to perform pretreatment susceptibility testing. But, when pretreatment susceptibility tests are not performed, it is important for the laboratory to freeze an initial isolate(s) and hold it for at least 6 months to permit baseline studies in the event the patient fails to respond to therapy. Drug susceptibility tests should be requested on subsequent iso- lates in the following instances: (1) For relapse or retreatment cases (2) When consideration is being given to change the drug regi- men because the patient's specimens: (a) Remain positive after 4 to 5 months on treatment (b) Become positive again after being negative (c) Show a sharp and persistent increase in acid-fast bacilli after an initial decrease (3) When primary drug resistance is suspected 4. Drug Resistance Drug resistance in Mycobacterium tuberculosis arises exclusively by spontaneous mutations that occur at random in the bacterial population and not by adaptation after exposure to the drug (6). A certain proportion of drug-resistant mutants exists in all populations of drug-susceptible tubercle bacilli. Fortunately, these mutants do not 163 survive well in normal environments, and although they arise at an estimated rate of about 10-8 per generation, they usually remain in the minority. But, when single drug therapy is used in the treatment of tuberculosis, the large population of susceptible organisms is inhib- ited or killed, thus leaving those few spontaneously occurring mutants (unaffected by the drug) to multiply until they comprise the majority of the bacterial population. This selective process is recognized as “ACQUIRED” RESISTANCE. The epidemiologic concept of PRIMARY DRUG RESISTANCE (PDR) refers to drug resistance in strains of tubercle bacilli isolated from patients who have never been treated with the antituberculosis drugs in question. These PDR cases were initially infected with drug-resistant bacteria excreted by a source case whose bacilli acquired resistance during treatment (3). Therefore, drug resistance in tuberculosis repre- sents an increase in the proportion of resistant cells resulting from an interplay of the natural phenomenon of spontaneous mutation and the selection to predominance of drug-resistant mutants by the action of single or ineffective drug therapy. This selective (acquired) resis- tance can occur only if bacterial mutation has already taken place before drug treatment is started. The CRITERIA of RESISTANCE in tuberculosis are based upon recognition of the fact that there is a certain proportion of drug resistant mutants above which therapeutic success is less likely to occur. This proportion has, with some statisti- cal measure of support, been set at 1% (4). When testing for drug resistance, one must be aware of the critical concentration of the drugs being tested. The critical concentration of a drug is the amount that inhibits the growth of most cells in wild strains of tubercle bacilli without appreciably affecting the growth of all mutants present (4). Table 10 gives the critical concentrations for drugs tested on Middlebrook 7H-10 agar in the laboratories of the Centers for Disease Control (7). TABLE 10. Critical concentrations of antituberculosis drugs in Middlebrook 7H-10 medium Drug Concentration in 7H-10 ( pg/ml) Isoniazid 0.2 Streptomycin 2.0 Ethambutol 5.0 Rifampin 1.0 p-Aminosalicylic acid 2.0 Ethionamide 5.0 Kanamycin 5.0 Capreomycin 10.0 D-Cycloserine 30.0 Pyrazinamide (at pH 5.5) 25.0 164 When 1% or more of the tested bacterial population becomes (or exhibits) resistant to the “critical” concentration of a given drug, that drug is not, or soon will not be, useful for continued anti-tuberculosis chemotherapy. 5. Methods There are three general methods used throughout the world for determining drug susceptibility of mycobacteria: the absolute con- centration method, the resistance ratio method, and the proportion method. When properly standardized and performed, all have been shown to provide clinically useful data. These three methods were described in detail in publications by the World Health Organization (3, 4) and in Tubercle (Leading Article, 1964; 45:169—-71). The absolute concentration method uses a standardized inoculum grown on control media and media containing appropriately graded concentrations of the drug(s) to be tested. Several concentrations of each drug are tested, and resistance is expressed in terms of the lowest concentration of the drug that inhibits all or almost all of the growth; i.e., minimal inhibitory concentration (MIC). This method is widely used in Middle and Eastern Europe. The resistance ratio method compares the resistance of unknown strains of tubercle bacilli with that of a standard laboratory strain. Parallel sets of media, containing twofold dilutions of the drug, are inoculated with a standard inoculum prepared from both the unknown and known (standard) strains of tubercle bacilli. Resistance is ex- pressed as the ratio of the minimal inhibitory concentration (MIC) of the test (unknown) strain divided by the MIC for the standard strain in the same set. This method was developed by the British primarily for testing susceptibility of M. tuberculosis to streptomycin. The proportion method enables precise quantitation of the propor- tion of mutants resistant to a given drug. Several dilutions (100-fold apart) of inoculum are planted in replicate onto both control and drug-containing media; at least one dilution should yield isolated countable (50 to 100) colonies. When these numbers are corrected by multiplying by the dilution of inoculum used, the total number of viable colonies observed on the control medium, and the number of mutant colonies resistant to the drug concentrations tested may be determined. The proportion of bacilli resistant to a given drug is then determined by comparing these numbers and expressing the resis- tant portion as a percentage of the total population tested. In every case the inoculum size must be known so the actual proportion of resistant mutants may be calculated. Most laboratories in the United States and France use this method. 165 6. Type of Susceptibility Test Although different media, such as L-J (7, 8) and 7H-11 (28), have been used in performing drug susceptibility tests, we strongly recom- mend the use of 7H-10 agar medium. The medium of choice is 7H-10 agar because of its simple composition and ease of preparation and because its transparency enables the earlier detection and more pre- cise quantitation of colonies. Because of the transparency of 7H-10 medium, a dissecting microscope often may be used to detect myco- bacterial growth before it is visible on opaque, egg-base media. Drug susceptibility tests may be performed by either the DIRECT or the INDIRECT method. For the direct method, the inoculum is a digested, decontaminated clinical specimen in which acid-fast bacilli may be demonstrated in stained smears. The inoculum size is adjusted on the basis of the numbers of bacilli observed in the smear, and the bacilli are representative of the bacillary population of a particular lesion(s) in the host. The direct test provides earlier test results (3 weeks). In contrast, the inoculum for the indirect test is a subculture of the organism previously grown on some primary isolation medium. The indirect test is used when (a) smears are negative but cultures positive, (b) growth on the control medium of a direct test is inadequate (less than 50 to 100 colonies), or (c) a reference culture is submitted for testing. The inoculum for the indirect test is adjusted turbidimetri- cally to yield isolated countable colonies on at least one of the dilu- tions (7). Because the bacterial inoculum for the indirect test is selected by the microbiologist, it is possible to select a proportion of suscepti- ble-to-resistant bacilli that does not reflect the true situation in the lesion(s) of the patient. For this reason the investigator is urged to pick a portion of all colonies growing on the culture. Details for performing each method follow: (1) DIRECT TEST (a) Stain and examine smears of concentrated clinical speci- mens. After observing the first acid-fast organism, record the average number of bacilli observed in the next 20 fields. Count clumps as only one organism, because they will give rise to only one colony on the medium. (b) Select dilutions of sputum concentrate to be used as inoc- ula on the basis of Ziehl-Neelsen smear results as follows: Microscopy Inoculate Less than 1 AFB/field Undiluted and 10-2 1-10 AFB/field 10" and 1073 More than 10 AFB/field 10-2 and 1074 166 (c) (d) (e) (f) (g) (h) (i) (i) (k) Make appropriate serial tenfold dilutions of the digested sputum concentrate. Use a capillary pipette to place three drops of inoculum onto each quadrant of the drug and control media. Inocu- late one set of plates with the higher dilution of inoculum and the duplicate plates with the lower dilution. If the patient has been under treatment, one plate should also be inoculated with undiluted inoculum, regardless of smear results. Bacilli from such patients may be stained, but because of drug action, many may be nonviable. Include a set of control plates inoculated with a drug- susceptible strain of M. tuberculosis (the H37Rv strain is commonly used). This serves as a quality control check on the presence of drug in the medium. Place plates, medium side down, in individual polyethyl- ene bags and seal. Incubate at 35 to 37°C in an atmosphere of 10% CO, (CO, not essential when performing indirect method). Read and report results of drug tests at 3 weeks. If cultures are fully matured, and ONLY IF THEY SHOW DRUG RESIS- TANCE, they may be reported in less than 3 weeks. Even though colonies may be fully matured on control media in less than 3 weeks, ‘drug susceptible’ reports should not be tendered until the third week, because resistant colo- nies often grow more slowly than susceptible ones and may not be visible until the third week. Colonies appearing on drug medium after 3 weeks do not necessarily repre- sent resistant bacilli. Some drugs are only bacteriostatic and, on prolonged incubation, may be inactivated to sub- static levels, permitting still fully susceptible colonies to grow after 3 weeks. Such colonies are usually smaller than those on the control quadrant. Do not discard as negative drug plates containing the con- trol quadrants until the end of the fifth or sixth week. This is not to permit a reading of the drug susceptibility test, but rather to use the control quadrant as another “tube” of isolation medium, because it has been inoculated with the digested-decontaminated specimen. Examine all grossly ‘negative’ quadrants with the aid of a dissecting microscope (30 x to 60 x); drug resistant or very slowly growing mycobacteria may be invisible to the naked eye. Record the colony characteristics on the control medium. Report results obtained with both dilutions of inoculum 167 and calculate the percentage of resistance. A convenient way to record growth is as follows: Confluent (500 or more colonies) 4+ Almost confluent (200 to 500 colonies) 3+ 100 to 200 colonies 2+ 50 to 100 colonies 1+ Less than 50 colonies Actual count Note: Designations like 2+ and 4+ do not permit precise quantitation (i.e., 2+ is not necessarily 50% of 4 +). Exact colony counts must be made on at least one control qua- drant, except as noted below. For a test to be valid, the control must show good growth (at least 50 to 100 colonies), but not so much as to simulate drug resistance due to growth of spontaneously occurring drug-resistant mutants (i.e., only confluent growth on control). In a properly performed drug susceptibility test, one of the control media should have countable colonies so that the percentage of resistant organisms can be pre- cisely calculated. The only time a control quadrant with confluent growth is acceptable as a valid test is when the culture exhibits total susceptibility to each drug tested. (2) INDIRECT TEST To prepare inoculum from growth on solid medium: (a) (b) (c) (d) (e) (f) Scrape several spadesful (2 to 5 mg) of growth from drug- free medium. Try to pick a portion from each colony. Transfer to a sterile 16 x 125-mm screwcap tube containing 6 to 8 glass or plastic beads and 3 ml of Tween-albumin liquid medium. Homogenize on a test tube mixer for 5 to 10 minutes. Allow larger particles to settle. Withdraw supernatant sus- pension and adjust density to that of a MacFarland No. 1* with sterile distilled water or saline. Dilute to 10 2 and 10 “ by making serial tenfold dilutions in sterile distilled water or saline. Proceed as for Direct Test (above) for inoculation, incuba- tion, and reading. To prepare inoculum with Tween-albumin liquid culture: (a) (b) Transfer a portion of each colony from drug-free medium into a Tween-albumin liquid medium, such as Middlebrook 7H-9. Incubate at 35°C for 7 days or until turbidity matches that of MacFarland No. 1 standard. * Prepare MacFarland No. 1 standard by adding 0.1 ml of 1% BaCl, to 9.9 ml of 1% H,SO,. 168 (c) Dilute to 10-3 and 1075 in sterile water or saline. (d) Inoculate the two dilutions, incubate, and read as described for the Direct Test (above). 7. Preparation of Drugs and Drug-Containing Media e Drug Potencies The true potency of the drug is the number of micro- grams of active drug per milligram total weight of the product. Not all antimicrobial drugs have been isolated in pure form, and a portion of their weight may be due to impurities or to the sulfate or other radical component of the molecule. The potency should be stated on the vial, otherwise it is necessary to consult the manufacturer for this information. Each lot of partially purified drugs will vary from previous ones, and the potency of one lot may not be the same as that of another lot. e Sterilization of Drugs Sterile lyophilized drug reconstituted with sterile distilled water (according to directions accompanying the vial) by using aseptic technique requires no further sterilization. Dilute with sterile water to obtain a working stock solution to add to the medium. For nonsterile powdered or crystalline drugs: Dissolve the drug in distilled water, or other solvent, to make a concentration of a known potency (e.g, 10,000 pg/ml). Sterilize by membrane filtration. Filtra- tion by Seitz or sintered glass causes loss of up to 30% potency, depending upon the nature of the drug and its concentration. Autoclave sterilization is not always acceptable because some drugs are inactivated by heat. For greater uniformity, the membrane filter is recommended. For filtration of small volumes, two membrane filter models are available: (a) microsyringe filter holder that attaches to a 10-ml syringe with Luer fittings and (b) microanalysis filter holder plus a 125-ml filtering flask. The microsyringe requires no vacuum source and is easily cleaned by rinsing in tap water, soaking overnight in distilled water, and drying in the air. A 25-mm filter, porosity 0.45 pm fits either of the “micro’’ models. A prefilter may be placed on top of the membrane filter. 169 For sterilization, assemble the filter holder with prefilter and filter paper in place. Autoclave 10 to 15 minutes at 121°C. Allow autoclave pressure to rise slowly and, at the end of the sterilization period, allow the autoclave to return slowly to normal so there will be no sudden pressure changes to damage the delicate filter paper. Allow filter to cool to room temperature before using. Note: Filtration of volumes less than 10 ml results in unnecessarily large loss of potency because the drug is absorbed during the early part of the filtration pro- cess. e Sources of Drugs When ordering drugs for laboratory purposes, the request should be directed to the medical department of the com- pany and not the sales department. Request information on the potency of the compound, since many of the antimi- crobial agents are commonly used as salts. Laboratory Division Dihydrostreptomycin or Charles Pfizer & Company, Inc. Streptomycin sulfate (SM) 235 East 42nd Street New York, NY 10017 Bristol Laboratories Kanamycin sulfate (KM) P.O. Box 657 Syracuse, NY 13201 The Lannett Company p-Aminosalicylic acid (PAS) 9000 State Road Philadelphia, PA 19136 Ives Laboratories Ethionamide (THA) 685 Third Street New York, NY 10017 Lilly Research Laboratories Capreomycin sulfate (CM) Indianapolis, IN 46206 Cycloserine (CS) Isoniazid (INH) Ciba Pharmaceutical Company Rifampin (RIF) Summit, NJ 07901 Aldrich Chemical Company Pyrazinamide (PZA) 940 W. Saint Paul Avenue Milwaukee, WI 53233 170 Lederle Laboratories Ethambuto/ (EMB) Pearl River, NY 10965 Sigma Chemical Company Streptomycin sulfate (SM) P.O. Box 14508 Cycloserine (CS) St. Louis, MO 63178 Isoniazid (INH) Ethionamide (THA) Ethambutol (EMB) Rifampin (RIF) Capreomycin sulfate (CM) Kanamycin sulfate (KM) Note: Store unopened vials of drug in the powder form according to directions on the label. Some require refriger- ation, others do not. Store opened vials of drug powder in a vacuum dessicator so they will not absorb moisture. To calculate the weight of drug necessary to prepare 10 ml of a 10,000 pg/ml solution, use this formula. 10,000 potency (in mg/g) X 10 = milligrams to weigh Example: Potency of dihydrostreptomycin sulfate = 800 mg of active SM per gram 10,000 _ —800 X10 = 125 Dissolve 125 mg in 10 ml of water for 10,000 ng SM per milliliter. e Preparation of Isoniazid (INH) Solution Potency = 1000 mg/g (1) Dissolve 100 mg (100,000 pg) of isoniazid in 10 ml of distilled water. This is a 10,000 pg/ml solution. (2) Sterilize by membrane filtration. (3) Add 1 ml of the above to 9 ml of sterile distilled water. This is a 1000 pg/ml solution. (4) Add 1 ml of 1000 ng/ml solution to 9 ml of sterile water. This is a 100 pg/ml solution. (5) Add 0.4 ml of 100 pg/ml solution to 200 ml of medium to obtain a final concentration of 0.2 ng INH per millili- ter of medium. (6) Add 2.0 ml of 100 pg/ml solution (or 0.2 ml of 1000 pg/ml solution) to 200 ml of medium if a final concen- tration of 1.0 ng INH per milliliter is to be used. 171 e Preparation of Streptomycin (SM) Solution If potency = 800 mg/g (1) (2) (3) (4) Dissolve 125 mg of dihydrostreptomycin sulfate in 10 ml of distilled water. This is a 10,000 ng/ml solution. Sterilize by membrane filtration. Add 1 ml of the above to 9 ml of sterile distilled water. This is a 1000 pg/ml solution. Add 0.4 ml of 1000 pg/ml solution to 200 ml of medium to obtain a final concentration of 2.0 ng SM per millili- ter of medium. e Preparation of PAS Solution If potency = 877.2 mg/g (1) (2) (3) (4) Dissolve 114 mg of sodium p-aminosalicylate in 10 ml of distilled water. This is a 10,000 ng/ml solution. Sterilize by membrane filtration. Add 1 ml of the above to 9 ml of sterile distilled water to obtain a 1000 wg/ml solution. Add 0.4 ml of 1000 ng/ml solution to 200 ml medium to obtain a final concentration of 2.0 wg PAS per millili- ter of medium. e Preparation of Kanamycin (KM) Solution If potency = 823 mg/g (1) (2) (3) (4) Dissolve 121.5 mg kanamycin sulfate in 10 ml of dis- tilled water. This is a 10,000 ng/ml solution. Sterilize by membrane filtration. Add 1 ml of the above to 9 ml of sterile distilled water. This is a 1000 pg/ml solution. Add 1.0 ml of 1000 ng/ml solution to 200 ml medium to obtain a final concentration of 5 ug of KM per millili- ter of medium. e Preparation of Ethionamide (THA) Solution If potency = 1000 mg/g (1) (2) (3) (4) Dissove 100 mg of ethionamide in 10 ml of dimethyl- sulfoxide (DMSO). This is a 10,000 ng/ml solution. Sterilize by membrane filtration. Add 1 ml of the above to 9 ml of sterile distilled water. This is a 1000 pg/ml solution. Add 1 ml of 1000 pg/ml to 200 ml solution medium to obtain a final concentration of 5 ug THA per milliliter of medium. e Preparation of Ethambutol (EMB) Solution Potency = 1000 mg/g (1) Dissolve 100 mg of d-ethambutol in 10 ml of distilled 172 water. This is a 10,000 ng/ml solution. (2) Sterilize by membrane filtration. (3) Add 1 ml of the above to 9 ml of sterile distilled water. This is a 1000 ng/ml solution. (4) Add 1.2 ml of 1000 pg/ml solution to 200 ml of medium to obtain a final concentration of 6 ng EMB per millili- ter of medium. Preparation of Cycloserine (CS) Solution Potency = 1000 mg/g (1) Dissolve 100 mg of cycloserine in 10 ml of distilled water. This gives a 10,000 ng/ml solution. (2) Sterilize by membrane filtration. (3) Add 0.6 ml of 10,000 ng/ml solution to 200 ml of medium to obtain a final concentration of 30 pg CS per milliliter of medium. Preparation of Rifampin (RIF) Solution Potency = 1000 mg/ml (1) Dissolve 100 mg of rifampin according to manufac- turer's instructions to make a 10,000 pg/ml solution. (2) Sterilize by membrane filtration. (3) Add 1 ml of the above to 9 ml of sterile distilled water. This is a 1000 pg/ml solution. (4) Add 0.2 ml of 1000 ng/ml solution to 200 ml of medium to obtain a final concentration of 1.0 pg of rifampin per milliliter of medium. Preparation of Capreomycin (CAP) Solution If potency = 839 mg/g (1) Dissolve 119.2 mg of capreomycin sulfate in 10 ml of distilled water. This gives a 10,000 pg/ml solution. (2) Sterilize by membrane filtration. (3) Add 1 ml of the above to 9 ml of sterile distilled water. This is a 1000 pg/ml solution. (4) Add 2.0 ml of 1000 pg/ml solution to 200 ml of medium to obtain a final concentration of 10.0 ng of capreomy- cin per milliliter of medium. Media Preparation Drug-containing media should be prepared only in quan- tities sufficient for immediate use because prolonged stor- age may affect drug potency. If medium is prepared in 200-m! amounts and poured into quadrant plates, each 200- ml batch will make 16 to 18 quadrants. Thus, to prepare one set of quadrant plates, prepare four flasks of enriched Middlebrook 7H-10 medium (three drug quadrants and one 173 control quadrant). Use 500-mI flasks for the medium. Fol- low directions on the bottle of commercially available Middlebrook 7H-10 agar medium base. To each 500-ml| flask, add the required amount of powdered base (for 200-ml total volume) and suspend in 180 ml of distilled water. Sterilize in the autoclave (121°C/10 minutes). Cool to 56°C in a water bath, and add 20 ml of oleic acid-albumin-dextrose- catalase (OADC) to each flask of medium. Make sure OADC is at room temperature before adding to agar. Add the proper amount of each desired drug (see table 11). Swirl to mix well (avoid bubbles) and dispense into sterile plates (5 ml per quadrant), using aseptic technique. Each plate should have a control quadrant without drug for accurate reading of the test results; example shown below. Standard Middlebrook 7H-10 base and oleic-albumin-dextrose- catalase (OADC) enrichment are used to prepare all drug- containing media, except for the medium used to test pyrazina- mide. The base of this medium is prepared by Gibco and buffered to pH 5.5 with albumin-dextrose enrichment being substituted for OADC (2a). 8. Storage of Drugs All antituberculosis drugs used in the preparation of susceptibility test media may be prepared in bulk from a given lot of drug. Correct for drug potency. Weigh drugs to the fourth decimal place on an analytical balance and solubilize in either water or methyl alcohol. TABLE 11. Preparation of drug media Drug Concentration of Volume of Drug Final Drug Drug Solution Solution (in ml) Concentration (ng/ml) Added to 200 ml (ng/ml Medium) Sterile 7H-10 Medium Isoniazid 100 0.4 0.2 Streptomycin 1000 0.4 2.0 Ethambutol 1000 1.2 6.0 Rifampin 1000 0.2 1.0 p-Aminosalicylic 1000 0.4 2.0 acid Ethionamide 1000 1.0 5.0 Kanamycin 1000 1.0 5.0 Capreomycin 1000 2.0 10.0 D-Cycloserine 10,000 0.6 30.0 Pyrazinamide 5000 1.0 25.0 (pH 5.5) 174 Sterilize the solution by membrane filtration (0.45 um pore size). Dis- pense these stock drug solutions into sterile 30-m| amber vials, seal, and store in a —70 to —80°C freezer until used. On the day the drug is to be added to the medium, remove from freezer, thaw to room temperature, and dilute with cooled sterile water, if needed. Discard excess solution and NEVER refreeze. The concentration of stock drug in the frozen vial and the volume that must be added to 200 ml of complete 7H-10 medium to attain the desired drug concentrations are shown in table 11. 9. Disc Method The submerged disc method of susceptibility testing first described in 1966 (42) has been critically evaluated (14, 15, 17) and shown to obviate many of the problems associated with susceptibility testing, such as instability of drugs, imprecision in weighing and diluting drugs, mislabeling of media, and the difficulty of preparing quality media. Commercially available paper discs containing appropriate concentrations of the primary drugs are listed in table 12. Not all antituberculosis drugs are available in disc form; if special requests are made for tests of susceptibility to drugs not listed in table 12, the laboratory must either prepare its own medium or refer the request to a reference laboratory where such tests are done (e.g., State health department, or even CDC, provided the request is routed through the State laboratory). TABLE 12. Drug-containing discs for susceptibility tests* Drug ng of Drug Final ng/ml in Disc** Drug in Medium Isoniazid 1 0.2 5 1.0 Streptomycin 10 2.0 50 10.0 Rifampin 5 1.0 25 5.0 Ethambutol 25 5.0 50 10.0 p-Aminosalicylic Acid 10 2.0 50 10.0 Ethionamide 25 5.0 Kanamycin 30 6.0 * Do not forget to include drug-free control quadrants in your susceptibility test. Remember: Duplicate sets of drug plates are needed for each test culture because two different inocula must be used. ** Discs are coded to identify both the drug and its concentration. 175 Appropriate drug discs are dispensed aseptically into the center of individual quadrants of sterile plastic dishes. This may be done while the basal medium is being sterilized in the autoclave or tempered at 50°C before the OADC enrichment is added. Exactly 5.0 ml of sterile, tempered (52°C), complete 7H-10 medium is pipetted over the disc. Remember, the disc must remain submerged and not be allowed to float to the top. Let the medium solidify. See figure 22 for example of drug susceptibility test media made by using commercially available drug-impregnated discs. Incubate the plates overnight at room temperature to permit the drug to diffuse uniformly. Discs containing ethambutol lose activity rapidly when allowed to diffuse overnight; after medium has solidi- fied over ethambutol discs, the plates should be placed at 5°C to permit diffusion of drug. e Storage of Disc Drug Media Since 7H-10 agar medium in plates dehydrates when stored, the 7H-10 drug medium should be used within 1 month of preparation. This recommendation is for drug medium prepared “from scratch’ with powdered drug. Because of the very low final concentrations of most antituberculosis drugs incorporated into media, coupled with the difficulty of weighing and diluting small amounts of drugs or their solutions, most investigators elect to make more-than-needed quantities of drug medium and store it for future use. This should be discouraged. FIGURE 22. Drug susceptibility test medium made with drug- impregnated discs 176 The proper use of drug-impregnated discs ensures both a sta- ble and precisely measured quantity of drug. The exact number of drug plates needed for a given day can be prepared without the worries of quantitation and stability; therefore, long-term storage is no longer a problem. This means that a drug medium can be prepared on the day before it is needed and problems of storage, dehydration, or loss of drug activity can be avoided. This alone should greatly improve the quality of drug susceptibil- ity test results. 10. Reporting Test Results The report should contain: The type of test, DIRECT or INDIRECT. The amount of growth on the control medium. The amount of growth on each drug medium. The concentration of each drug in the medium. A rough calculation of the percentage of bacilli resistant to the drug. This formula may be used to calculate the percentage of resistant cells: Number of colonies on the drug . X 100 = % resistant Number of colonies on the control ° EXAMPLE: Test a Growth on Drug Undilute 10°? % Resistant Control 4+ 150C INH 0.2 pg 1+ 9C 6 SM 2.0 ng 0 0 0 EMB 5.0 ng 0 0 0 9 colonies X 100 = 6% 150 colonies Inthis example, 6% of the organisms were resistantto 0.2 pg/ml of INH. 177 EXAMPLE: Test b Growth on Drug 10 103 % Resistant Control 4+ 60C INH 0.2 ng 120C 0 2 SM 2.0 pg 0 0 0 RIF 1.0 ng 0 0 0 The 10" control inoculum gave 4+, or colonies too numerous to count. The hundred fold dilution (10-3) gave 60 countable colonies. Therefore, the 4+ = 60 x 100 or 6000 colonies. Knowing this figure, the proportion of resistant bacilli may be calculated as 2% to 0.2 pg/ml INH. 120 —_ — 929 5000 X 100 = 2% An alternative way to calculate the percentage of resistant bacilli in Example Test b above is the following. Because the 10-2 dilution represents 1/100th (or 1%) of the 10" dilution, any colonies growing on drug medium inoculated with the 10" dilution that equal (or exceed) the number of colonies growing on the control medium inoc- ulated with the 10-3 dilution represent 1%, or more, of the test population. Thus, to quickly calculate the percentage of resistant cells, use this formula: Colony count on drug medium at lower dilution (10~" in Example) “” = % resistance Colony count on control medium at higher dilution (1073 in Example) In Example Test b above, 120 = 2% resistance. 60 If more than 1% of the test population is resistant to the drug under test, this suggests that resistance to that drug has developed or is in an advanced stage of development. 178 INH 0.2 SM 2.0 The inoculum (107) on the right shows 4 growth on the controls and INH quadrants, 23 colonies on SM, and 2 on PAS. A 100-fold dilution (103) of the inoculum on a second drug plate shows 59 colonies on the control, 54 on INH, and none on SM and PAS. Hence, 54/59 gives 92% resistance to INH. To determine percent resistant to SM and PAS, multiply 59 X 100 = 5900 colony units inoculated on to each quadrant of the 10! plate. 23/5900 X 100 — 0.4% R to SM. 2/5900 X 100 = 0.03% R to PAS. Ne NN] (Zip Cod 3 First (| Repeat I T— i; Mo. Year Laboratory Report ExamngE MICROSCOPICALLY BY: ZIEHL-NEELSEN FLUOROCHROME ACID FAST BACILLI: [Jrouna [JNot Founda Ee ow CO numerous FUNGI OR YEAST: Found Oot Found a Split Given to Mycology Lab. DRUG SUSCEBXIBILITY TESTS IN 7H-10 AGAR sd [JinoirecT Drug Growth * at Dils. Percent (pgm) wl CONTROL 0.0 100 INH 0.2 INH 1.0 2.0 10.0 2.0 FINAL REPORT MAY REQUIRE 8 WEEKS 5.0 5.0 SPECIMEN UNSATISFACTORY FOR TESTING BECAUSE: Od Insufficient Amount [| Leaked In Transit Not mycobacterium Oa Contaminated Oa 5.0 cs 20.0 ETH 5.0 *4 += >500,3+=200— 500, 2+=100—200,1 + =50 —100 <50 recorded & followed by ‘'c'*, as 10C = 10 colonies TENTATIVE IDENTIFICATION: IDENTIFICATION . tuberculosis [I] M. bovis . kansasii . marinum . scrofulaceum . szulgai Od M. gordonae Om. [2] M. avium complex (intracellulare) O M. xenopi Cm. a M. triviale J M. gastri OJ M. fortuitum complex a Rapid Grower [J other Mycobacterium flavescens terrae complex Day Date Received and Laboratory Specimen Number Month Year o Date Reported 1st 2nd Form No. Specimen Number Form Control Number SAMPLE LABORATORY REPORT FORM FIGURE 23. Drug susceptibility test 179 INH 0.2 SM 2.0 The inoculum on the plate at the right (1077) is too heavy to determine actual colony counts. In the hundredfold dilution of inoculum (103) on the plate to the left we count 41 colonies on the control and 17 on the PAS quadrant. Hence, the apparent resistance to PAS observed in the heavily inoculated plate is a true one, and approximately 41% of the population was resistant to PAS (17/41 X 100 = 41%). ~T (Zip Code) 0 Pst Mo. Day Year 0 Repeat Laboratory Report DRUG SUSCEPTIBILITY TESTS IN 7H-10 AGAR | IDENTIFICATION EXAMINGD MICROSCOPICALLY pIRECT [J INDIRECT CY suiberesi . tuberculosis BY: ZIEHL-NEELSEN Dion Growth* at Dils. Parcant O M. bovis FLUOROCHROME (Hg/m1) oo 0 " , = . kansasii ACID FAST BACILLI: CONTROL 0.0 id 3 m. marinum Foss [J Not Founda INH 0.2 Bn . scrofulaceum R, NH 10 MO M. szulgaei ew 20 a M. gordonae Numerous FUNGI OR YEAST: 10% CIM. iavescens dO Found Cnet Found 20 Oa M. avium complex O Split Given to Mycology Lab. 10.0 (intracellulare) FINAL REPORT MAY 5.0 [J m. xenopi REQUIRE 8 WEEKS 5.0 0 TT —_m FR TESTING BECAUSE. 2 Elm. crivite Od Insufficient Amount 22 Ol M. gastri ETH 5.0 Od Leaked In Transit C3 Not mycobacterium a Contaminated J M. fortuitum complex *4 + =>500,3+=200—500, TENTATIVE 24+ =100—200,1+=50—100 'DENTIFICATION: [3 Rapid Grower <50 recorded & followed by [J other Mycobactarium [| *c"”, as 10C = 10 colonies Date Received and Laboratory Specimen Number Date Reported Form Month Day Year Form No. Specimen Number 1st 2nd 3rd Sopra SAMPLE LABORATORY REPORT FORM FIGURE 24. Drug susceptibility test 180 11. Radiometric Methods Several rapid radiometric methods for susceptibility testing of mycobacteria have been reported recently (1, 2, 21, 27, 29, 30, 38, 40). Middlebrook and associates (30) used a medium containing radiola- beled palmitate as a carbon source. Growth of M. tuberculosis was detected by measuring the conversion of the substrate to “CO, in an ionization chamber. This technique has been the most widely accepted of the radiometric methods and when the procedure was compared to standard growth susceptibility tests, agreement appeared good in laboratories with a high degree of expertise in mycobacteriology, whereas results varied in those laboratories where proficiency was not as well-maintained. Refinements in the methodology are being made as more and more laboratories investigate the application of this rapid technique. As in any new technique, the users must criti- cally control the procedure against a known standard method to famil- iarize themselves with the new methodology and some of the poten- tial pitfalls. This implies a thorough familiarity with, and capability to conduct, parallel control studies using conventional methodology to ensure that results are comparable before making a dramatic shift from conventional to radiometric methodology. Early in the study of this rapid technique it was observed that the radiometric method had poor correlation with conventional methods when tubercle bacilli were resistant to drugs, but these problems are gradually being resolved and we should soon see wider use of the new methodology. While contemplating the use of radiometric methods, a laboratory should determine: (a) if the shorter test period is justification for the greater expense and (b) if earlier drug susceptibility reports contribute signifi- cantly to patient care (since most smear-positive patients are started on treatment before the report is issued). The step from use of radiometric methodology for drug susceptibil- ity testing to its use for primary isolation of mycobacteria from clini- cal specimens is not a difficult one, and one should expect increasing activities in this area in the coming years. 12. Drug Assay The assay of antituberculosis drugs in serum or urine was once recommended for the management of the individual case of tubercu- losis (36). Today, serum assays are most often performed in special cases when there is reason to believe the drug is not being absorbed or eliminated properly by the patient (e.g., as a result of intestinal tract surgery (33) or kidney disease (37) or to monitor levels of drugs known to exhibit a narrow margin between therapeutic success and toxicity (e.g., cycloserine and some of the aminoglycosides in some patients) (28). There are many methods, both biological and chemical, 181 for determining serum levels of most antituberculosis drugs. Some biological assay methods for isoniazid, ethionamide, ethambutol, and rifampin utilize tube dilutions of the patient's serum to inhibit growth of a test strain of bacteria (36, 41), whereas other biological assay methods employ a serum-soaked paper disc diffusion technique to detect serum drug levels of streptomycin, capreomycin, and kanamy- cin (36, 37). There are chemical assay methods for detecting p-amino- salicylic acid (26), pyrazinamide (35) and cycloserine (20, 35). Several simple and reliable drug-screening tests have been described for detecting antituberculosis drugs or their metabolites in urine (5, 10, 11, 24, 35, 39). The need to determine drug levels in most patients is quite uncom- mon. On rare occasions however, it might be beneficial to help evalu- ate patient progress. In such cases, consult some general references (5, 25, 28) and then contact a Level lll reference laboratory that special- izes in this procedure. REFERENCES 1. Arroyo J, Medoff G. Mycobacterium chelonei infection: successful treatment based on a radiometric susceptibility test. Antimicrob Agents Chemother 1977;11:763-4. la. Bailey WC, Bass JB, Hawkins JE, Kubica GP, Wallace RJ Jr. Drug susceptibility testing for mycobacteria. ATS News, Winter 1984:9-10. 2. Benitez P, Medoff G, Kobayashi GS. Rapid radiometric method of testing suscepti- bility of mycobacteria and slow growing fungi to antimicrobial agents. Antimicrob Ag Chemother 1974;6:29-33. 2a. Butler WR, Kilburn JO. Improved method for testing susceptibility of Mycobacterium tuberculosis to pyrazinamide. J Clin Microbiol 1982;16: 1106-9. 3. Canetti G, Froman S, Grosset J, Hauduroy P, Langerova M, Mahler HT, Meissner G, Mitchison DA, Sula L. Mycobacteria: laboratory methods for testing drug sensitiv- ity and resistance. Bull WHO 1963;29:565-78. 4. Canetti G, Rist N, Grosset J. Mesure de la sensibilité du bacille tuberculeux aux drogues antibacillaires par la méthode des proportions. Rev Tuberc 1963;27: 217-72. 5. Chadwick MV. Mycobacteria. Institute of Medical Laboratory Sciences Monographs. FJ Baker ed Boston: Wright, 1982. 6. David HL. Probability distribution of drug-resistant mutants in unselected popula- tions of Mycobacterium tuberculosis. Appl Microbiol 1970;20: 810-4. 7. David HL. Fundamentals of drug susceptibility testing in tuberculosis, Atlanta: Centers for Disease Control, PHS, HEW, 1971 (HEW publication no. 00-2165). 8. David HL. The bacteriology of the mycobacterioses. Atlanta: Centers for Disease Control, PHS, HEW, 1975 (HEW publication no. 76-8316). 9. Demerec M. Origin of bacterial resistance to antibiotics. J Bacteriol 1948;56:63. 10. Eidus L, Hamilton EJ. A new method for the determination of acetylisoniazid in the urine of ambulatory patients. Am Rev Respir Dis 1964;80:587-8. 11. Eidus L, Harnavansingh AMT. A urine test for control of ingestion of ethionamide. Am Rev Respir Dis 1968;98:315-6. 182 12. 14. 15. 16. 17. 18. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. Farer LS. All about TB: what the practicing physician must know and can do about tuberculosis. Clin Notes Respir Dis 1978;16:3. . Ferebee SH, Palmer CE. Prevention of experimental tuberculosis with isoniazid. Am Rev Tuberc 1956;73:1. Griffith M, Barrett HL, Bodily HL, Wood RM. Drug susceptibility tests for tuberculo- sis using drug impregnated discs. Am J Clin Pathol 1967;47: 812-7. Griffith ME, Matajack ML, Bissett ML, Wood RM. Comparative field test of drug- impregnated discs for susceptibility testing of mycobacteria. Am Rev Respir Dis 1971;103:423-6. Grosset J. Bacteriologic basis of short-course chemotherapy for tuberculosis. Clinics in Chest Med 1980;1:231-41. Hawkins JE. Disc and dilution media for susceptibility testing against primary antituberculosis drugs (abstract). Am Rev Respir Dis 1976;113/4 (part 2):78. Hawkins JE, Good RC, Kubica GP, Gangadharam PR, Gruft H, Stottmeier K. Ameri- can Thoracic Society policy statement on levels of laboratory services for myco- bacterial diseases. Am Rev Respir Dis 1983;128:213. . Hoeprich PD. Chemoprophlylaxis in infectious disease. In: Hoeprick PD, ed. Infec- tious Diseases. New York: Harper and Row, 1972:207. Jones CR. Colorimetric determination of cycloserine, a new antibiotic. Anal Chem 1956,28:39-41. Kertcher JA, Chen MF, Charache P, Hwangbo C, Camargo EE, Mcintyre PA, Wagner HN Jr. Rapid radiometric susceptibility testing of Mycobacterium tuberculosis. Am Rev Respir Dis 1978;117:631-7. Kubica GP. Susceptibility testing of tubercle bacilli. In: Bondi A, Bartola JT, Prier JE, eds. The clinical laboratory as an aid in chemotherapy of infectious disease. Baltimore: University Park Press, 1977:107-14. Kubica GP, Dye WE. Laboratory methods for clinical and public health mycobacteri- ology. Atlanta: Centers for Disease Control, PHS, HEW, 1967. Lederle Laboratories, Medical Communication Department. Myambutol, ethambu- tol hydrochloride, 2nd ed. 1968:68-9. Lorian V. Antibiotics and chemotherapeutic agents in clinical and laboratory practice. Springfield: Charles C. Thomas, 1966. Marshall EK Jr. Determination of p-aminosalicylic acid in blood. Proc Soc Exp Biol Med 1948; 68:471-2. McClatchy JK. Rapid method of microbial susceptibility testing. Infect Immun 1970;1:421-2. McClatchy JK. Antituberculosis drugs: mechanisms of action, drug resistance, susceptibility testing and assays of activity in biological fluids. In: Lorian V. ed. Antibiotics in laboratory medicine. Baltimore: Williams and Wilkins, 1980:135-69. McDaniel RE, Abensohn MK, Spoon DR, Kobayashi GS, Medoff G, Marr JJ. A rapid radiometric method for determining the sensitivity of clinical isolates of Mycobac- terium tuberculosis to several chemotherapeutic agents. J Lab Clin Med 1977;89: 861-7. Middlebrook G, Reggiardo Z, Tigertt WD. Automatable radiometric detection of growth of Mycobacterium tuberculosis in selective media. Am Rev Respir Dis 1977;115:1066-9. Mitchison DA. Basic mechanisms of chemotherapy. Chest 1979;76: 771-80. Mitchison DA, Dickinson JM. Bactericidal mechanisms in short-term chemother- apy of tuberculosis. Bull Int Union Tuberc 1978;53:270. Pickleman JR, Evans LS, Kane JM, et al. Tuberculosis after jejunoileal by pass for obesity. JAMA 1975;234:744. Pyle M. Relative numbers of resistant tubercle bacilli in sputa of patients before and during treatment with streptomycin. Proc Mayo Clinic 1947,22:465. Rao KVN, Eidus L, Jacob CV, Tripathy SP. A simple test for detection of pyrazinamide and cycloserine in urine. Tubercle 1965;46:199-205. 183 36. 37. 38. 39. 40. 41. 42. Russell WF Jr, Middlebrook G. Chemotherapy of tuberculosis. Springfield, Ill: Charles C Thomas, 1961. Sabath LD. A simple rapid microassay for nephrotoxic antibiotics. Scope Mono- graph. Kalamazoo: Upjohn Company, 1972. Siddiqi SH, Libonati JP, Middlebrook G. Evaluation of a rapid radiometric method for drug susceptibility testing of Mycobacterium tuberculosis. J Clin Microbiol 1981;13:908-12. Simpson J. Simple tests for the detection of urinary PAS. Tubercle 1956;37:33-4. Snider DE Jr, Good RC, Kilburn JO, et al. Rapid drug-susceptibility testing of Mycobacterium tuberculosis. Am Rev Respir Dis 1981;123:402-6. Stottmeier KD, Woodley CL, Kubica GP, Beam RE. A simple biological method for determination of small amounts of tuberculostatic agents in fluids. Bull WHO 1967;37:961-6. Wayne LG, Krasnow |. Preparation of tuberculosis susceptibility testing mediums by means of impregnated discs. Am J Clin Pathol 1966;45:769-71. 184 Reporting Culture Results Reporting Culture Results Personnel in the mycobacteriology laboratory must establish a uni- form procedure for examining and recording culture observations. By using some type of magnification (a hand lens or a dissecting microscope), the microbiologist can observe cultural growth earlier, can detect transparent colony forms more easily, and can note detailed morphology of mature colonies. The culture report should reflect the following observations: Growth Rate: The number of days required for colonies to appear Temperature: The temperature at which growth occurred Pigment Production: The mycobacteria may be placed into three large groups on the basis of pigment formation (Identification Test Techniques page 104). Colony Morphology: Information obtained from observation of col- ony morphology often proves helpful in the identification of cul- tured organisms For best observation of colony morphology, the microbial suspen- sion inoculated onto media must be diluted sufficiently to yield iso- lated colonies. The following are some terms used to describe myco- bacterial colonies on culture media. All workers in a given laboratory should agree on the meaning of these terms as they apply to the different colony forms of mycobacteria. Examples of these various terms are given in color plates 23 to 37 on pages 147 to 157. Surface: This may be smooth (SM), rough (R), or corded, or “X" colonies. The latter are dense, compact, smooth-to rough, slow- growing colonies that often exhibit tiny stick-like projections at the periphery when viewed by transmitted light. Because the colo- nies are almost unique to M. xenopi, they are referred to as “X" colonies (the letter coming from the species name). Elevation and Colony Form: Descriptive terms include flat, convex, pyramidal, umbonate (raised center), rosette (lobate), doughnut. Edge: This may be entire, irregular, flat, and spreading. Opacity: Colonies may be described as transparent, translucent, or opaque. Pigment (in light and dark): Record this as nonpigmented (buff), pigmented (name the color, e.g., yellow, orange), or “Greening” (some cultures absorb the malachite green dye from L-J slants during incubation, or when they are stored in the refrigerator). 185 Two additional terms commonly used in the mycobacteriology lab- oratory to refer to culture growth are “eugonic” and “dysgonic”. The term “eugonic” describes cultural growth that is luxuriant (i.e., large, easily seen colonies), whereas ““dysgonic” refers to tiny, difficult-to- see colonies that might be said to be growing poorly. Quantity of Growth — Record as Follows: No colonies No AFB grown <50 colonies Actual count 50-100 colonies 1+ 100-200 colonies = 2+ 200-500 colonies = 3+ (almost confluent) Over 500 colonies = 4+ (confluent) If laboratory findings are to be useful, they must be communi- cated to the proper authorities. Clinicians use the findings in the diagnosis and management of disease; public health authorities use the findings for statistical and epidemiologic purposes, and to control that bacteriologically proven cases are receiving appropriate therapy. Laboratory procedures for mycobacteriology are notoriously time- consuming, often taking weeks or even months to complete. For this reason, interim reports should be issued. They may be sent out after (a) acid-fast smears have been examined, (b) growth has been observed on the culture medium and the species has been tentatively identified or results of a direct drug susceptibility test are known, and finally, (c) the species has been precisely identified on the basis of detailed in vitro tests and/or indirect drug susceptibility tests have been completed. Obviously, all of the technical details of each laboratory test cannot be recorded on the reports, but they should be precisely outlined in the laboratory procedure manual, and the results must be a part of the daily laboratory records. The validity of these results also must be insured by including appropriate quality controls with each batch of media or each test run. The interim reports may be provided to the clinician by mailing copies of the original report form as new information becomes available, or by using precarboned, sectioned, perforated forms. In the latter case, the appropriate section of the form may be completed, dated, detached, and mailed. The final report will have all the data previously reported, so earlier interim reports can be destroyed, and only the final report is to be retained in the patient's file. A copy of the final report (and perhaps even the interim reports) should also be sent to the public health authorities. 186 Thus, smear reports should be completed and returned to the clini- cian within a few days of receipt of specimen; contaminated cultures may be noted within the first 7 to 10 days and repeat specimens may be requested; preliminary (tentative) identification and/or direct drug susceptibility test results should be available within 3 to 4 weeks; all test results may not be available for 6 to 9 weeks. A sample report form is shown on this page. LABORATORY NAME & ADDRESS FORM NAME & NUMBER Patient Name I [> [ Address City State Zip Code Social Security Number Please Return Report To Physician's Code To be Filled in by Physician Day | Year Date Specimen Collected Or. SPECIMEN Patient on Therapy? (Yes (No Sputum Service Requested: 0 Natural 0 Mx 0 culture O Drugs O Induced 0 ine O pas 0 0 Gastric Osm OETA 0 a Urine 0 ems OO kM a O Other Orr O a (Specify) If repeat, date |Mo. Day |Year O Culture of last specimen INTERIM REPORT #1 ACID FAST BACILLI Fungi or Yeast SPECIMEN UNSATISFACTORY EXAMINED MICROSCOPICALLY| [] found [J Not found OJ fount 0 Nottouny |FORTESTING BECAUSE By [J Ziehl-Neelsen 0) 1.2 per 300 fields O split sent to Mycology [J insufficient Amount 3 Fluorochrome 0 1.9/100 fields [11-9/10 fields Qu bacteria hort [J Leaked in transit 0 19/tield 0 >9/field ound ot found pies OJ split to Gen'l. Bact'y. [J Contaminated (Send Another) INTERIM REPORT #2 REPORT #3 DRUG SUSCEPTIBILITY TESTS IN 7H-10 AGAR 0 FINAL REPORT [J Tentative Report 0 Direct (J Indirect Growth *® at dil'n Percent of [J Isolate specifically identified as Drug ug/ml Pop’In. Mycobacterium tuberculosis CONTROL 100 INH 0.2 INH 1.0 [J Culture sent to Reference Laboratory SM 20 Final report may take 4-6 weeks SM 10.0 7 + entative identification or rough grouping suggests RIF 1.0 RIF 50 [1 M. tuberculosis EMB 5.0 [J Photochromogen [J Scotochromogen PAS 2.0 ETA 50 |] omogen [J Rapid Grower KM 5.0 — *4+ = >500, 3+ = 200-500, 2+ = 100-200, 1+ = 50 colonies, Date reports mailed <50 recorded, followed by “"c”’, as 10c = 10 colonies. 1st 2d 3d If >1% of tested population exhibits resistance to Date specimen rec'd Lab. specimen No given drug, this suggests emergence of drug resistance. (For Training Use Only) 187 Quality Control in the Mycobacteriology Laboratory Quality Control in the Mycobacteriology Laboratory Quality control procedures should be performed on a regualr basis in the mycobacteriology laboratory to assure reproducibility and reli- ability of laboratory results. The importance of laboratory test results in the practice of medicine and the increasing complexity of many modern laboratory procedures make it essential that good quality control measures be instituted to monitor the rapidly expanding, often automated, laboratory technology. For a quality control program to be helpful, it must be practical and workable. Quality control is the responsibility of all laboratory personnel, but if a full-time person is assigned to ‘‘quality control,” that person should be qualified in microbiology. When applicable, quality control procedures must meet the require- ments of the Clinical Laboratories Improvement Act of 1967. Today, the majority of clinical laboratories in the United States are under the jurisdiction of one or more accreditation agencies, such as Medicare, the College of American Pathologists, the Joint Commission on the Accreditation of Hospitals, and a number of State and local public health agencies. A well-designed and properly managed quality control program is an asset to any laboratory. Some of the positive aspects of a quality control program follow: Potential problems in the isolation and identification of microorga- nisms can be greatly reduced by monitoring media and reagents before using them on clinical specimens. Serious and costly breakdown of equipment may be minimized by routine monitoring and maintenance. Laboratory reports can be more expeditious and accurate because the use of poor or inadequate media, equipment, and techniques is minimized. The quality control program can also be a learning technique ena- bling the recognition or identification of problem areas that might otherwise have been overlooked. The keys to the development of a workable quality control program are (a) adequately trained, interested and committed personnel and (b) common-sense use of practical procedures. 189 A. General Recommendations 1. 2. All quality control records should be retained in files or notebooks for at least 2 years. Procedural manual(s) should be available for every routine procedure performed in the laboratory. The microbiologist in charge should date and initial all changes in techniques. All containers of media, stains, and reagents should show the date received and the date first opened. Each material should be periodically rechecked, and if found to be un- satisfactory, it should be removed immediately from the laboratory. Purchases should be limited to a 6 months’ supply. Laboratory procedures used routinely should be those that have been published in reputable microbiological books, manuals, or journals. Procedures that have not been de- scribed in publications may be used, provided the neces- sary experimental control studies have been performed in a competent manner and a description of the procedure is expected to be published. Laboratories should maintain the number and variety of cultures needed to check the quality of tests performed. B. Laboratory Arrangement and Personnel 1. Supplies, equipment, and work area should be arranged to facilitate an efficient work flow. Work areas should be kept free of dust. Benches should be swabbed at least once a day with an acceptable disinfectant. Personnel employed in the mycobateriology laboratory should be tuberculin tested and X-rayed, as suggested in the “Safety in the Laboratory’ chapter. C. Laboratory Equipment Equipment should meet the manufacturer's claims and specifica- tions in the user's laboratory. The user should monitor equipment regularly to assure the constant accuracy and precision necessary for quality laboratory performance. Table 13 indicates the monitoring necessary for equipment found in the mycobacteriology laboratory. 190 TABLE 13. Monitoring of Equipment Item Monitor Procedure Autoclave Record temperature each run; hold record in files. Re- cording thermometer is advisable. Use cultural device (spore strips, spore suspension) monthly; more frequently, if there is evidence of con- tamination. Decontaminate waste materials for 1 hour. Biological Safety Airflow checks: Cabinet (BSC) Use anemometer to measure rate of airflow across front Biological Safety Cabinet (BSC) opening; should be 75 linear feet/minute for Class | BSC; 75 to 100 linear feet/minute in Class Il BSC, depending on type (check with manufacturer). Install a magnehelic gauge in the exhaust duct to mea- sure any pressure drop across the bacterial filter and mark the limits when the airflow across the front opening is not optimal. Use smoke sticks to determine the pattern of air currents in the room, and especially when new equipment is placed in the BSC work space or when any change is made in the room ventilation pattern. Filters: Replace when gauge in exhaust duct indicates that airflow across front opening has dropped below optimal levels. CAUTION: Entire BSC must be decontaminated before filters are removed. Germicidal ultraviolet (UV) lights: Use UV light meter to measure light intensity after initial burn of 100 hours. Test every 6 months. Replace lamp when reading drops to 70% of the inital reading. CAU- TION: Clean UV lamps every 2 weeks with alcohol-soaked cotton. Do not purchase UV lamps far in advance of need. UV lamps are commonly included in Class | BSC, but only on special request in most Class Il Safety Cabinets. The value of these lamps in a well-maintained, properly moni- tored BSC is questionable. Careful cleaning and swab- bing of BSC work surfaces with an effective disinfectant is probably more effective. UV can be very effective in control of airborne mycobacteria that escape the confines of the BSC. Installation of such fixtures should be done only after consultation with personnel knowledgeable in their use. 191 TABLE 13. Monitoring of Equipment—Continued Item Monitor Procedure Centrifuge Incubator, 35°C CO, Microscope pH meter Water baths Refrigerator 2 to 8°C Freezers 0 to —70°C Glassware Use tachometer. Use appropriate rpm on an 8- to 12-place horizontal or angular head centrifuge to attain centrifugal force of 3000 x g for sedimentation of mycobacteria. CAUTION: A 16-place horizontal head usually cannot achieve this cen- trifugal force. The use of lesser g-forces for longer periods may prove lethal to mycobacteria because of heat build- up during spinning. Check brushes and bearings every 6 months. Use recording thermometer, if available. If not available, record temperature daily, preferably in a.m. Test temper- ature at several sites within incubator; thermometer should be placed in water reservoir (e.g., Erlenmeyer flask). Control light by covering glass front of incubator door and restricting use of any lights inside the incubator. Check CO; content daily with CO, indicator, such as Fyrite. Always clean oil immersion objective after examining a positive acid-fast smear. Clean microscope after each use. Keep microscope under dust cover when not in use. Compensate for temperature with each run. Date buffer solutions and discard when unsatisfactory. Standardize with pH 4.0 and 7.0 buffers before each test or series of tests. Check temperature before and during use. Clean monthly. Check temperature daily. Walk-in should have recording thermometer. Connect walk-in to alarm system. Clean monthly. Defrost or check refrigerator and freezer com- partment every 3 months. Check daily. Connect to alarm system. Clean every 6 months. Discard chipped or etched glassware. Glassware should be free of detergents. Check sterilized glassware for sterility. Do not store sterile glassware for more than 3 weeks before it is used. Use oil-free aluminum foil to cap glassware before it is sterilized (do not use paper). 192 TABLE 13. Monitoring of Equipment—Continued Item Monitor Procedure Culture Purity Check by acid-fast staining and/or inoculating plated media to obtain isolated colonies at each transfer or sub- culture. Stains Quality control stains by preparing smears of a known acid-fast positive sputum or tissue sample (M. tubercu- losis). Use new clean slides and AFB-free water. In addition to the in-house monitoring of laboratory personnel, space, and equipment, it is important that other quality checks be made, beginning with the specimen and ending with the final report. eo Specimen It is important to note the quality of the sputum; i.e., is it thick, mucoid, purulent, or is it more like saliva? The latter is important, primarily when a negative sputum smear report is sent. Also impor- tant for the negative smear or culture report is the quantity of speci- men submitted; if the quantity is <5 ml and the report is negative, notify the physician who submitted the specimen(s). e Transportation Best results are obtained with specimens that are transported quickly (<7 days) to the laboratory. Any delays in delivery should be noted on the report form, particularly with negative reports. e Microscopy A number of checks should be done in this area. Each new batch of stains should be checked both on known positive and known nega- tive control smears; the use of positive control ensures the staining capability of the new stain solutions, whereas the use of negative control smears will confirm that acid-fast contaminants are not pres- ent in the stain solution(s) (i.e., from the water or reagents used to prepare the stains). It is well to confirm positive smears of clinical specimens by a second reader and to retain positive smears for sev- eral weeks or months against the likelihood of a request to “re- examine’ the smear. Records should be kept, perhaps even plotted on a graph, of numbers (or percentages) of positive smears observed for each time period (weekly, monthly, etc.); any sharp deviations from the norm should be investigated. Specimens that are smear- positive are generally culture-positive, unless the patient is on therapy; smear-positive, culture-negative specimens occur rarely (generally, less than 2%). 193 e Culture Media Records should be kept on all “homemade’’ media: source and batch number of reagents or powdered base media; coagulation time and temperature for inspissated egg media; visual appearance (e.g., color, bubbles, consistency); and firmness of media. Records should be kept of numbers or percentages of specimens positive by culture over predetermined periods (weekly, monthly). Because of the greater sensitivity of culture, only about half of culture-positive specimens will also be smear-positive, but nearly all smear-positive specimens should be culture-positive (if the medium is well made). All prepared or commercially purchased media should be checked for sterility and sensitivity. The latter may be effected by using a standardized suspen- sion of M. tuberculosis (e.g., H37Ra or R1Rv available from ATCC) that has been checked for viable count and stored at —70°C. If it is known that this suspension may be diluted (e.g., to 107°) and that platings of 0.1 ml will yield known colony numbers (e.g., 100 += 50), all “good” batches of new media should fall in this range; inferior quality media would commonly not support growth of colonies at as high a dilution (e.g., perhaps only at 10-4 or 10-3). e Digestion-Decontamination To determine decontaminating capabilities of each new batch of reagent, digest 4 to 6 sputa, concentrate by centrifugation and inocu- late to plates of “general bacteriology agar media’ (e.g., blood, brain heart infusion, MacConkey) as well as “TB media” (L-J, 7H-10 or 7H-11). Numbers of contaminants that grow after 24 to 48 hours at 37°C should be minimal-to-none. Records should be kept of the per- centage of clinical specimens contaminated; acceptable range is 3% to 5%. Contamination significantly less than 3% suggests overly harsh decontamination, whereas percentages much greater than 5% sug- gests either (a) too weak a decontaminant or (b) incomplete digestion. Additionally, a careful recording of the numbers (percentage) of iso- lates of M. gordonae can serve as a built-in indicator of digestant toxicity. This species is commonly seen in most laboratories (>10% of routine isolation) and is more susceptible to toxic digestants than is M. tuberculosis, so recovery of fewer than 5% M. gordonae (from among all AFB cultured from clinical specimens) suggests an overly harsh decontamination action. eo Water, Distilled and Tap Both should be checked periodically for presence of acid-fast con- taminants and should be the first thing considered when untoward results are encountered in the laboratory (e.g., excessive numbers of 194 smear-positive, culture-negative specimens; ‘““mini-epidemics” of isolation of saprophytic mycobacteria, such as M. gordonae or M. terrae complex). If water appears cloudy or dirty, simple centrifuga- tion at 3000 xg (or greater) of 200 to 250 ml of water (multiple tubes, if necessary) may sediment enough material to enable detection of acid-fast bacilli by smear examination. Alternatively, one may (a) use the Candida method to facilitate sedimentation (1) or (b) pass one liter of water through a sterile 0.22 um pore size membrane filter and either (a) place the intact filter on the surface of a plate of 7H-10 or 7H-11 agar or (b) cut the filter into strips (sterile scissors) and asepti- cally place onto the slants of L-J egg medium. Manuals of methods and daily records used in the laboratory must be maintained and kept current. D. Media, Reagents, and Biochemical Tests Table 14 indicates the minimum number of tests for the quality control of media, reagents, and biochemical tests. These control tests should be performed each time fresh media or reagents are made, and each time tests are run, except as otherwise noted. Quality control procedures in table 14 provide checks for Sterility, Growth, and Biochemical Response. Sterility tests should be per- formed at 25°C and 37°C on a sample of each batch of medium that is autoclaved or filter-sterilized during preparation. Sterile media or sub- strates are inoculated at the same time and in the same manner as regular tests, except that “‘sterile’’ reagents or diluents are used as inoculum. This is a control for “environmental” contaminants. Growth studies verify that the medium will support growth of the desired organism(s), whereas positive- and negative-control organisms are used to document that the medium will produce the expected bio- chemical test responses. 195 961L TABLE 14. Quality Control Procedures (2,6) Biochemical Medium or Reagents Sterility Growth Response Organism(s) Expected Results Arysulfatase, M. smegmatis, TMC 1515, Negative (no 3-day or M. avium complex, color change) X X TMC 1403 M. fortuitum, Positive (pink to TMC 1529 red). Compare to color standards. Catalase, M. avium complex, < 45mm Semiquantitative TMC 1403 M. gordonae TMC 1324, or >45 mm X X M. kansasii, TMC 1201 Catalase, M. gastri, TMC 1456, or Negative pH 7.0, 68°C M. tuberculosis, (No bubbles) X TMC 201 M. fortuitum, TMC 1529, Positive or M. gordonae, (bubbles) TMC 1324 may take 5 minutes after reagent added L6l TABLE 14. Quality Control Procedures (2,6) (Cont.) Medium or Reagents Biochemical Organism(s) Expected Results Iron Uptake M. chelonae, Negative TMC 1544 (no color change or growth) M. fortuitum, Positive TMC 1529 (Rusty brown growth) Nitrate Reduction M. chelonae, TMC 1544 Negative (no color change after reagents but red after zinc added) M. tuberculosis, 3+ tob+ TMC 201 (pink to deep red after reagents) Pyrazinamidase M. bovis BCG, Negative (no pink TMC 1011 band) M. avium complex, Positive (pink TMC 1403 band) Sodium Chloride M. bovis BCG, Negative (no growth) Tolerance TMC 1011 M. fortuitum, TMC 1529 Positive (growth) 861 TABLE 14. Quality Control Procedures (2,6) (Cont.) Medium or Reagents Biochemical Organism(s) Expected Results MacConkey Agar Without Crystal M. phlei, TMC 1548, or M. bovis BCG, TMC 1011 Negative (no growth in 11 days) Violet ; ” M. fortuitum, Positive (growth TMC 1529 in 5 days with or without color change) Niacin M. fortuitum, Negative (no color Standard or Strip TMC 1529 change) Test (If 7H-10 used must use warm (37°C) extracting fluid and incubate media at 37°C for 2 hr. to get good niacin extraction) M. tuberculosis, TMC 201 Positive (yellow color) must have 50-100 colonies on 3- to 4-week-old culture Tellurite Reduction (3-day) M. triviale, TMC 1453 Negative (no black precipitate) M. avium complex, TMC 1403 Positive (black precipitate) 661 TABLE 14. Quality Control Procedures (2,6) (Cont.) Biochemical Medium or Reagents Sterility Growth Response Thiophen-2- Carboxylic Acid Hydrazide (TCH) Susceptibility X X Tween 80 Hydrolysis Organism(s) M. bovis BCG, TMC 1011 Expected Results Negative (no growth) M. tuberculosis, TMC 201 M. avium, complex, Positive (growth) Negative (no change X X TMC 1403 in 10 days) M. kansasii, Positive (pink-to- TMC 1201 red) Tween Opacity M. avium, complex, Negative (no band) TMC 1403 X X Ls M. flavescens, Positive (opaque TMC 1541 band formed in 1 week) Urease M. avium complex, Negative (no color TMC 1403 change) X X M. scrofulaceum, Positive (pink to TMC 1302, or M. chelonae, TMC 1544 red) 00¢ TABLE 14 . Quality Control Procedures (2,6) (Cont.) Biochemical Medium or Reagents Sterility Growth Response Organism(s) Expected Results Photochromogencity Test provides own negative control X X X M. kansasii, Positive (color TMC 1201 change after 2-hr. light exposure) Caps must be loose during light exposure. Growth on Isolation Check pH, Media (L-J and color and 7H-10) consistency M. tuberculosis, H37Ra Should be able to support growth of small inoculum in X X 4 to 6 weeks Media for Drug M. tuberculosis, known Positive on Susceptibility susceptible strain control; Testing X X TMC 201 or TMC 102 inhibited by all drugs. Drug-free medium inoculated with test culture Verifies ability of the medium to support growth of test strain. Procedure for the Storage of Mycobacterial Stock Cultures 1. 2. 3. Place 2 ml of tap water in a 2-dram, flat-bottom, screwcap vial. Cap the vials loosely and autoclave them at 121°C for 15 minutes. Prepare suspensions of the cultures by emulsifying five or six colonies in the 2 ml of water. Alternatively, the cultures may be grown in 7H-9 broth and bot- tled in 1- to 2-ml amounts in the above vials or in vaccine-stoppered vials (4,5). Cap the vials tightly and store upright in a freezer at a temperature ranging from —16 to —70°C. Some species (especially M. tubercu- losis) lose viability on prolonged storage (1 year or more) at —20°C, but all species retain virtually 100% viability when stored at —70°C (5). Note: The contents of the vials may be thawed and refrozen sev- eral times without appreciable loss in viable count. Sources of Mycobacterial Stock Cultures 1. 2. American Type Culture Collection (ATCC) 12301 Parklawn Drive. Rockville, MD 20852 Your State health department laboratory Standard Strains for Quality Control A minimal number of ‘standard’’ strains of mycobacteria that may be used to control the quality of media and the positive and negative reactions of most commonly used biochemical tests is listed below. M. avium, arylsulfatase, 3-day (=) TMC 1403 pyrazinamidase (+) semiquantitative catalase (<45) tellurite reduction (+) Tween hydrolysis (=) Tween opacity (=) urease (=) M. bovis, BCG heat stable catalase (=) TMC 1011 MacConkey agar (=) pyrazinamidase (=) sodium chloride tolerance (—) thiophen-2-carboxylic acid hydrazide, growth on (—) M. chelonae, iron uptake (=) TMC 1544 nitrate reduction (=) urease (+) M. flavescens, Tween opacity (+) TMC 1541 201 M. fortuitum, arylsulfatase, 3-day (+) TMC 1529 heat stable catalase (+) iron uptake (+) MacConkey (+) niacin {—} sodium chloride tolerance (+) M. kansasii, MacConkey (=) TMC 1201 photochromogenic (+) semiquantitative catalase (>45) Tween hydrolysis {+) M. triviale, tellurite reduction (—) TMC 1453 sodium chloride tolerance (+) M. tuberculosis, heat stable catalase (—) R1Rv or TMC 205 niacin (+) nitrate reduction (+) thiophen-2-carboxylic acid hydrazide, growth on (+) M. tuberculosis, H37Ra or TMC 201 growth on egg and agar media; drug test REFERENCES 1. Dizon D, Milhailescu C, Bae HC. Simple procedure for detection of Mycobacterium gordonae in water causing false-positive acid-fast smears. J Clin Microbiol 1976;3: 211. 2. Ellis RJ. Manual of quality control procedures for microbiological laboratories. Atlanta: Center for Disease Control, PHS, HEW, 1974. 3. Jones WD Jr. Simple method for maintaining stock cultures of Mycobacterium species. Am J Clin Pathol 1957;27:363—4. 4. Kim TH, Kubica GP. Long term preservation and storage of mycobacteria, Appl Microbiol 1972;24:311-7. 5. Kim TH, Kubica GP. Preservation of mycobacteria: 100% viability of suspensions stored at —70°C. Appl Microbiol 1973;25:956 60. 6. Miller JM. Quality control in the microbiology laboratory. Atlanta: Centers for Dis- ease Control, PHS, HEW, 1983. 202 Appendix Equipment and Supplies Required in a Mycobacteriology Laboratory The items listed may be obtained from most general laboratory supply companies. General Equipment Autoclave, steam or electric Balances, top loading (analytical, if needed for drugs) Beads, glass or plastic Burner, Bunsen or Fisher Carbon dioxide gas Carbon dioxide gauge, Fyrite or Burwell Centrifuge with tachometer, 3000 x g Desiccator, vacuum (for drug storage) Discard cans, splash proof Discard pans with covers Filters, bacteriological (Seitz and Millipore) and vacuum pump or line for sterilization of drugs, albumin, etc. Freezer: —20 to —70°C Hand lens, 10X Incubators, fitted for carbon dioxide Inoculating spade or loop Inspissator (for homemade egg medium, if used) Interval timers Lamps or light with magnifier (3X — 10X) for culture examination Mailing containers, double for specimen and culture shipment. Microscope, dissecting, 10—50X for use with transmitted light Microscope for fluorescence microscopy Microscope with oil immersion lens Mixer, test tube, high speed; e.g., Vortex or other pH meter Pipetting unit, Cornwall for dispensing media Racks for specimen tubes Refrigerator, 4 to 10°C Safety equipment (see below) Slide staining rack, electric or adjustable sink top rods Slide warmer Stirrer, magnetic 203 Temperature block heater 37 and 68°C Thermometers Turntable for inoculating plates Water baths, 37 and 68°C, and if needed, 56°C to inactivate albumin Glassware Bottles for stains Cylinders, graduated, 25 ml to 1000 ml Flasks, Erlenmeyer or other, 50 ml to 3000 ml Flasks, Volumetric, 100 ml, 500 ml, 1000 ml Pipettes Capillary (Pasteur) 1 ml graduated in 1/100 ml 5 ml graduated in 1/10 ml 10 ml graduated in 1/10 ml Slides, microscope 27 x 75 mm frosted end, regular 27 x 75 mm for fluorescence microscopy Tubes, screwcap 13 x 100 mm 16 x 125 mm 20 x 150 mm Plastics Bags, mylar (CO,-impermeable) Bags, polyethylene Centrifuge tubes, screwcap, 50-ml, water-tight Petri dishes Plain 15 x 100-mm bi-plate (2 sections) 15 x 100-mm X-plate (4 sections) 15 x 100-mm Safety Alcohol-sand flask Airflow-testing device Meters Smoke sticks Autoclave testing device Thermometer, recording Sterility-testing indicator Biological Safety Cabinet (BSC) Centrifuge safety cups, aerosol free Discard pans with covers 204 Disinfectant for hands, 2% amphyl, 70% alcohol or other Disinfectant, liquid phenolic or other tuberculocidal agent Filters, absolute (HEPA, or high efficiency particulate air) Fogging machine (if needed for room with recirculating air) Protective clothing: Caps Gloves Gowns Masks Shoe covers Safety pipetting devices Splash-proof containers Tissue grinder, Ten Broeck, hand Ultraviolet lamps in BSC (if present in original equipment) Ultraviolet intensity meter Chemicals, Media Bases, and Miscellaneous Supplies Only chemicals of the highest purity should be used. A few may be available from only one manufacturer. Chemicals Acridine Orange Agar Albumin, Bovine Fraction V Alcohol, 95% ethyl Ammonium hydroxide Ammonium sulfate Aniline Asparagine Auramine O Barium chloride Biotin Bromthymol blue Calcium chloride -2H,0 Catalase, technical grade Copper sulfate -5H,0 Cyanogen bromide Dextrose (glucose) Disodium phosphate -12H,0 and/or anhydrous Ferric ammonium citrate (green) Ferrous ammonium sulfate Fuchsin, basic, 94% dye content Glycerol, reagent grade Hydrochloric acid 205 Hydrogen peroxide (30% Superoxol) Magnesium citrate Magnesium sulfate -7H,0 Malachite green oxlate Methylene blue chloride Monopotassium phosphate (anhydrous) Monosodium glutamate N-acetyl-L-cysteine powder Neutral red N-naphthylethylenediamine dihydrochloride Oleic acid Phenol crystals, reagent grade Phenol liquid Phenolphthalein Phenolphthalein disulfate, tripotassium salt Phenolsulfonphthalein, sodium salt (phenol red) Potassium permanganate Potassium tellurite Potato flour or potato starch Pyrazinamide Pyridoxine hydrochloride Pyruvic acid, sodium salt Sodium carbonate Sodium chloride Sodium citrate -2H,0 Sodium hydroxide Sodium nitrate Sulfuric acid 2-thiophene carboxylic acid hydrazide (TCH) Trisodium phosphate Tween 80 Zephiran (17% benzalkonium chloride) Zinc metal powder, (dust) Zinc sulfate -7H,0 Media Bases Dubos broth base with ADC enrichment Lowenstein-Jensen powdered egg base MacConkey agar (no crystal violet) Middlebrook 7H-9 broth base with ADC enrichment Middlebrook 7H-10 agar base with OADC enrichment Urease agar base, Difco-Bacto Wayne Sulfatase agar 206 Miscellaneous Supplies Niacin test strips Nitrate test crystalline reagents (if used) Urea discs 207 3372723128523 116 GENERAL LIBRARY - U.C. BERKELEY BO0OOLS5LOS0