Recent advances in laboratory procedures for pathogenic mycobacteria

Clin Lab Med 23 (2003) 801–821

Recent advances in laboratory procedures for pathogenic mycobacteria Robert C. Cooksey, PhD Tuberculosis/Mycobacteriology Branch, Centers for Disease Control and Prevention, Mail Stop F08, 1600 Clifton Road, Atlanta, GA 30333, USA

Mycobacteriology is an area in which technological advances are highly warranted, not only because of the clinical importance of members of the genus Mycobacterium but also because of their diversity, fastidiousness, and generally slow growth characteristics. The most notable species in this genus is Mycobacterium tuberculosis, but many other species also are primary pathogens [1–4]. Isolation of mycobacteria has traditionally been performed through the use of selective in vitro growth media. Conventional identification is usually pursued by biochemical testing and growth characteristics, and antimicrobial susceptibility testing is based on growth in the presence of the drug of interest [5,6]. Laboratory diagnosis of Mycobacterium tuberculosis The most important components of the traditional presumptive diagnosis of tuberculosis are clinical symptoms and chest X ray [5]. Laboratory confirmation of a presumptive diagnosis includes acid fast bacillus (AFB) smear microscopy followed by culture [6]. Although positive AFB microscopy results are generally indicative for the presence of members of the genus Mycobacterium, they are not specific for M tuberculosis. The sensitivity of AFB smear microscopy using the traditional Ziehl-Neelsen staining method ranges from 22% to 78% compared with culture results, and the limit of detection is approximately 5  103 to 1  104 bacilli/mL, depending on several variables. These factors include the type of specimen, the Mycobacterium species present, the efficiency of decontamination, liquefaction and concentration of the specimen, and technical preparation of the slide and expertise of the laboratory personnel performing the microscopic examination [7]. E-mail address: [email protected] 0272-2712/03/$ - see front matter Ó 2003 Elsevier Inc. All rights reserved. doi:10.1016/S0272-2712(03)00085-4

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Whereas the appearance of visible growth after inoculating processed sputum onto conventional solid media (eg, Lowenstein-Jensen medium or Middlebrook 7H10 or 7H11 agar) typically requires 3 to 4 weeks, newer culture methods are less dependent on their ability to visualize actual growth; rather, they are more dependent on measurements of metabolic products and can significantly reduce time to detection of growth. The BACTEC 460 System (Becton-Dickinson Microbiology Systems, Sparks, Maryland) was the first semi-automated instrument dedicated to this purpose [8]. The principle, reported in 1977 by Middlebrook et al [9], is based on the measurement of 14CO2 released from 14C-palmitic acid during cellular metabolism. Cultures are sampled periodically using a needle that pierces rubber septums on the broth culture vials; growth indices commensurate with bacterial growth on a scale from 0 to 999 are determined. The BACTEC 960 System has been introduced by Becton-Dickinson as an alternative to—and perhaps replacement for—the BACTEC 460 System [10]. The basic principle of this instrument is the same as for the Mycobacterial Growth Indicator Tube (MGIT), which has an oxygensensitive quenching compound imbedded in silicon at the bottom of tubes containing liquid culture medium supplemented with a mycobacterial growth enhancer (oleic acid-albumin dextrose) and an antimicrobial cocktail to inhibit contaminants [11]. Fluorescence may be manually observed following illumination with a 365 nm UV lamp or quantitated in the BACTEC 960 System instrument if the growth of an organism consumes dissolved oxygen. The BACTEC MGIT 960 System enables detection of the growth of mycobacteria starting with a clinical specimen in approximately 2 weeks and offers recovery rates that are comparable with the BACTEC 460 System instrument [12]. The MB/BacT System (Organon Teknika, Inc., Durham, North Carolina) is a liquid culture system that uses automated colorimetric monitoring of CO2 in a closed system [12]. A solid-state sensor located at the base of each culture vial contains a colorimetric indicator that changes from green to yellow when CO2 is produced within the vial [12]. Rohner et al [13] reported that times for recovery of mycobacteria averaged 17.5 days for the MB/BacT System, compared with 14.3 days for the BACTEC 460 System instrument in an evaluation of 73 specimens from which mycobacteria were recovered. Recovery rates and times that are comparable to the BACTEC 460 System have been reported for the ESP Culture System II (Trek Diagnostic Systems, West Lake, Ohio), which uses a unique principle of automated monitoring of pressure changes inside tubes due to production or consumption of gas during growth of organisms. In the early 1990s, Becton-Dickinson introduced Septi-Chek, a convenient system for the recovery of mycobacteria that is based on visible growth. The essential component of this recovery system is a 20-mL 7H9 broth tube containing antimicrobial agents to suppress growth of nonmycobacterial organisms, an atmosphere containing CO2, and a paddle containing three different agars. Although recovery times are longer using Septi-Chek

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compared with the BACTEC systems, the rates of recovery of mycobacteria were found to be comparable to the BACTEC 460 System [14]. Clinical mycobacteriology laboratories do not typically report negative cultures from any culture method for 6 weeks or more due to the possible presence of unusual dysgonic mycobacterial organisms. Two genotypic methods for detecting and identifying M tuberculosis bacilli and other species that are classically included in the M tuberculosis complex (MTC; M bovis, M africanum, and M microti) in patient specimens have been approved by the United States Food and Drug Administration (FDA) for clinical diagnostic use in the United States and offer promise for reducing turnaround times for laboratory diagnosis of tuberculosis [7]. Both methods are based on nucleic acid amplification. The Enhanced Amplified Mycobacterium Tuberculosis Direct Test (E-MTD; Gen-Probe, Inc., San Diego, California) has been approved for use on respiratory specimens regardless of AFB results. The Amplicor MTB assay (Roche Molecular Systems, Branchburg, New Jersey) is approved for use only with AFBpositive specimens. The Amplicor Assay is based on amplification of a genus-specific region of the 16S rRNA gene; after amplification, the products are denatured and hybridized to oligonucleotide probes specific for MTC organisms bound to wells of microdilution plates; and detection is achieved through the use of a bound biotin-avidin-horseradish peroxidase complex [15]. The E-MTD assay is based on reverse transcription of MTCspecific rRNA leading ultimately to the production of double-stranded cDNA, which is transcribed by DNA-directed RNA polymerase to produce additional rRNA template molecules in a cyclic fashion. These RNA amplicons are detected by a hybridization protection assay using an acridinium ester-labeled MTC-specific DNA probe. Studies that included evaluations of the E-MTD and Amplicor MTB assays were recently reviewed by Roberts et al [12] and Woods [7], who concluded that both assays are useful for the rapid diagnosis of tuberculosis. Becton-Dickinson introduced an alternative semi-automated kit for detecting MTC organisms in respiratory specimens in 1998 which was based on isothermal strand displacement amplification (SDA) [16,17]. The most recent version of this kit, the BDProbeTec ET, couples the SDA reaction to enhanced detection using fluorescence resonant energy transfer (FRET) chemistry, and although it has not received approval from the United States FDA for use in the United States, it is currently available in international markets. The sensitivity for detecting MTC in AFB-positive smear samples using BDProbeTec ET was found to be high (93.8%) in a recent evaluation [16]. Conventional antituberculosis drug susceptibility testing methods Laboratories were urged by the Centers for Disease Control and Prevention in 1993 to provide results of susceptibility testing with first-line drugs of M tuberculosis isolates within 30 days [18]. The goals of in vitro

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antimicrobial susceptibility testing of M tuberculosis should include not only the accurate detection of resistance in the shortest possible time, but also the identification of subpopulations of resistant bacilli that may compromise therapeutic success of antituberculosis drugs [5]. The two most frequently used methods in the United States for susceptibility testing of M tuberculosis isolates are agar proportion [19] and BACTEC [20]. When performing the agar proportion method, at least two dilutions of the test isolate are plated onto agar (eg, Middlebrook Complete 7H-10 or -11 agar) that contain concentrations of drugs considered to be ‘‘critical’’ as well as drug-free control agar. The critical concentration is the amount of drug that inhibits the growth of most cells in wild strains of tubercle bacilli without appreciably affecting the growth of all mutants present [5]. A digested and concentrated (ie, ‘‘processed’’) specimen, diluted according to the number of bacilli observed during acid-fast microscopic examination, may also be used as the inoculum in performance of the direct susceptibility test. The agar plates are incubated for 3 weeks, colonies are counted, and resistance is indicated if the number of colonies on the drug-containing agar is 1% or greater of the count on the control agar. Although this method does permit at least semiquantitation of resistant subpopulations and direct observation of contaminants, it is expensive in terms of time, as well as cost of media and some antimicrobial agents. Although the method overcomes problems associated with degradations of drug and media, novel drugs and alternative media should be evaluated for this potential problem. An alternative to preparing drug stocks is placing commercial antibiotic disks in petri dishes before pouring agar [8,21]. The BACTEC 460 System is an alternative to the agar proportion method for susceptibility testing of M tuberculosis, especially for clinical laboratories processing larger numbers of isolates [8]. Inocula are prepared by adjusting the turbidity of homogenized cell suspensions to that of a MacFarland no. 1 standard. Drug vials are inoculated with 0.1 mL of these suspensions and 0.1 mL of 1:100 dilutions of the suspensions are used to inoculate drug-free control vials. Results are usually available 4 to 7 days after inoculation and an organism is judged to be susceptible to a given drug if a change in the growth index for the control vial is greater than that for the drug-containing vial. Conversely, the instrument’s presentation of a growth index in a drug vial that is equal to or greater than that in the control vial indicates resistance. The media is commercially prepackaged (excluding antimicrobial agents) and contains polyoxyethylene stearate to enhance the growth of poorly growing mycobacterial isolates and to minimize cell clumping. The broth vials are inoculated and sampled semiautomatically with needles located on the instrument. Unlike the BACTEC 460 System, the more recently introduced BACTEC MGIT 960 System poses no hazards associated with the use of 14C-labeled substrates or needles. Inocula may be prepared from either liquid or solid media and susceptibility results are automatically interpreted by the instrument. The BACTEC 960 System has been evaluated, and the methods

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standardized for the primary drugs including rifampin (RIF), isoniazid (INH), ethambutol (EMB), streptomycin (STR), and pyrazinamide (PZA) [22,23]. It received approval by the US FDA for in vitro susceptibility testing of these drugs against M tuberculosis in the United States in 2002. The average turnaround time for M tuberculosis susceptibility results using the MGIT system ranges from 4.6 to 11.7 days after inoculation compared with 4 to 10 days for the BACTEC 460 System [22]. The ESP Culture System II (ESPII) and MB/BacT System, which were described above for isolating M tuberculosis organisms from specimens, also have been useful for antituberculosis susceptibility testing by reducing turnaround times for results and have been shown to have good agreement overall with results of more established methods [24,25]. Among rapid drug susceptibility methods for MTC, the National Committee for Clinical Laboratory Standards only recommends tests that have received US FDA clearance; these tests currently include the ESPII, the BACTEC 460 System, and the BACTEC 960 System [26]. A colorimetric assay for determining antituberculosis drug susceptibilities was described in 1995 by Yajko et al [27], who used an oxidationreduction dye, Alamar Blue (resazurin), as a growth indicator. Growth and metabolism of M tuberculosis in standard complete Middlebrook 7H9 broth was indicated by a color change of the dye from blue to pink, which is visually detectable without additional instrumentation. Although the Alamar Blue assay is insensitive to the presence of resistant subpopulations compared with the agar proportion or BACTEC method, it has been shown to be useful in research laboratories for determination of minimum inhibitory concentration (MICs) of standard antituberculosis drugs in 14 days or less after inoculation. It should be noted that clinical mycobacteriology laboratories do not determine in vitro drug MICs. Alamar Blue was adapted for performance in microdilution plates (microplate Alamar Blue assay; MABA) and parameters, including media composition, reaction time and temperature and inoculum preparation were standardized to achieve MIC results comparable to critical concentration drug susceptibility results obtained using the BACTEC 460 system in 1997 [28]. However, the Alamar Blue susceptibility assay has not received FDA approval. Another method for susceptibility testing of mycobacteria that does not involve expensive instrumentation is the Epsilometer test (Etest; AB Biodisk, Solna, Sweden) [29–31]. Antimicrobic gradients are prepared on plastic strips that are placed in contact with seeded agar surfaces; results for slowly growing mycobacteria are available 5 to 10 days after inoculation from actively growing broth cultures. Inocula prepared from highly turbid cultures (equivalent to 1.0 MacFarland turbidity standard) may increase the risks associated with infectious aerosols. The method offers MIC testing without advanced preparation of antimicrobic-containing media. Colonies growing inside zones of inhibition around the gradient strips also may reveal information about the presence of contaminants as well as subpopulations

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of bacilli with more than one level of resistance. In a recent evaluation of 95 isolates (40 of which were resistant to 1 or more primary drugs), agreement betweeen the Etest and agar proportion methods was 97.9% for RIF, 96.0% for EMB, 94.7% for INH, and 85.3% for STR [32]. Bacteriophages have also been exploited to measure the in vitro susceptibility of tubercle bacilli to antimicrobial agents. A recombinant mycobacteriophage (phAE40) into which a luciferase reporter gene (LUC) under control of a powerful promoter (hsp60) was cloned was used to infect M tuberculosis isolates that had been exposed to drugs. Bacilli that were inhibited by the drugs were incapable of supporting phage replication or gene expression, and measurable production of photons by the luciferase was diminished [33]. Light signals were detectable among strains infected with the recombinant phage and among mutants that were resistant to INH, STR, or RIF, whereas no light was detectable for susceptible strains exposed to these drugs [33]. More recently, phage D29 was used in the development of the phage amplified biologically (PhaB) assay, which is currently available commercially as the FASTPlaqueTB Assay (Biotec Laboratories Ltd., Suffolk, United Kingdom). Briefly, tubercle bacilli in processed sputum samples are infected with the phage, which is sequestered from a virucidal agent. After the virucide is neutralized, a phage-susceptible indicator organism is applied using agar plate overlays and plaques form if mycobacterial organisms are present in the specimen. The original phage infection is blocked by antimicrobial agents in susceptible strains [34,35], and susceptibilities to RIF, PZA, EMB, INH, STR, and ciprofloxacin were recently evaluated among 157 isolates . Results were available 2 to 3 days after initial exposure of the organism to drugs, and correlation with results of a conventional phenotypic resistance method ranged from 87% (PZA) to 100% (CIP) [34]. Mycobacterial esterases have also been exploited in the search for in vitro susceptibility testing methods. These enzymes can cleave fluorescein diacetate and derivatives such as sulfofluorescein diacetate to yield fluorescein, which may be excited followed by the emission of 485 nm fluorescence. Antimicrobial agents that inhibit or kill bacilli cause reduced esterase activity and a concomitant loss of fluorescence. A flow cytometric method that employs this principle and does not require multiplication of organisms in culture was described by Norden et al in 1995 [36]. The method, however, has not been extensively evaluated, and potential contamination of flow cytometers with viable bacilli may pose additional risks of infection unless the instruments are properly housed and used in appropriate containment facilities. Genotypic antituberculosis drug resistance testing Identification of drug resistance at the genetic level (eg, by using DNA probes) was proposed many years ago [37], and amplification assays,

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principally through polymerase chain reaction (PCR), have facilitated this approach [38–40]. Five points regarding the limitations of genetic detection of resistance should be considered: first, mutations in genes associated with antimicrobial resistance do not always equate with expression of phenotypic drug resistance; second, because most drug resistance in slowly growing mycobacteria is caused by point mutations, nucleic acid probe and amplification methods are of limited value; third, mutations for each primary drug are acquired independently of the others and therefore identifying a mutation associated with resistance for one drug cannot be used to presume resistance to another; fourth, resistance genes in mycobacteria may share sequences with nonmycobacterial genera and nonspecific PCR products may be obtained unless the purity of the mycobacterial DNA is confirmed; and fifth, some genotypic methods may be incapable of identifying mutations present as subpopulations among an abundance of wild-type alleles. DNA sequencing offers perhaps the most reliable method for detecting mutations. By using an automated sequencer, multiple isolates can be analyzed simultaneously and very high accuracy can be obtained, especially if both strands of duplex DNA are sequenced independently. Sequencing templates are prepared by PCR, and crude DNAs are usually suitable for this purpose. Although sequencing offers the additional advantage of detecting all mutations in a template, including those that have not been recognized previously, the association of phenotypic resistance with the novel mutation must be confirmed. Priming regions for sequencing are most often internal to those chosen for preparation of the sequencing template, and multiple, perhaps overlapping regions may need sequencing if important mutations for a particular drug are found throughout a large region ([500 bp). Among primary antituberculosis drugs, PZA resistance requires the most extensive search for mutations in a single gene. The product of pncA (pyrazinamidase) converts PZA to its active form, pyrazinoic acid. Mutations that affect either the expression or function of this enzyme may occur throughout the structural gene open reading frame (561 bp) or in regulatory regions located upstream [41]. Morlock et al [42] sequenced sense and antisense strands of 2 overlapping regions (4 sequencing reactions for each template) to analyze 744-bp pncA regions of 37 PZA-resistant strains. Mutations were found in 34 isolates (91.8%) and occurred from base ÿ16 to base 511. For other primary drugs such as isoniazid, the search for relevant mutations may be more narrowly focused. Mutations in codon 315 of the catalase gene, the enzyme of which is responsible for activating INH in the tubercle cell (katG), have been reported in 34% [43] to 94% [44,45] of INHresistant strains depending on the collection of isolates under investigation, but the overall prevalence is believed to be approximately 80% [46,47]. Screening isolates for mutations in the codon 315 region of katG using sequence analysis or alternative mutation identification methods is therefore

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straightforward relative to the search for important pncA mutations. But unlike PZA, for which only a single gene as been implicated in resistance, genes other than katG may play roles in INH resistance, including inhA, kasA, and ahpC [47,48]. Mutations within a single gene are most often associated with resistance to each of the three remaining primary antituberculosis drugs. These include RIF, for which mutations in an 81-bp (‘‘hotspot’’) region of the beta subunit of the gene encoding DNA-dependent RNA polymerase (rpoB) [49] have been found in approximately 95% of resistant strains [50–52]. For EMB, mutations in embB, which is a component of multigene operon involved in synthesis of a drug target enzyme arabinosyltransferase, were found in approximately 69% of EMB-resistant strains by Sreevatsan et al [53]. The majority of these (89%) were found at a single codon (no. 306). For STR, high-level resistance (500 lg/ml) is associated with mutations in rpsL, which encodes the drug’s target, S12 ribosomal protein [54–56], a mechanism that has been reported in other bacterial genera [47,57]. Although low-level STR resistance remains largely unexplained, some association with mutations in rrs that affect two loops of 16S rRNA may be involved in some strains [56,58]. Although DNA sequencing is the most definitive method of identifying mutations associated with drug resistance, hardware and expendables are expensive. Furthermore, mutations that are present as subpopulations may not be recognized by automatic base–calling software. Alternative genotypic methods to identify the most prevalent mutations have been described [46– 48,59–62]. Many of the methods are DNA probe–based, and one such method, the INNO-LiPA Rif.TB line probe assay for RIF resistance (Innogenetics N.V., Zwijndrecht, Belgium), has been offered as a commercial kit in countries outside the United States. The kit contains oligonucleodtides immobilized on nitrocellulose strips; one is specific for identifying the MTC, five are for the wild-type sequences for the 81-bp rpoB hotspot, and four are for the most frequently reported rpoB mutations. Test DNA is amplified with biotinylated primers, the products are hybridized with the immobilized probes, and hybrids are detected colorimetrically. A susceptible M tuberculosis strain results in the appearance of colored bands for the MTC probe as well as for each of the 5 rpoB wild-type probes. A strain with one of the four mutations will be negative for one of the wild-type probes and positive for the corresponding mutation probe [63]. In a comparison with DNA sequence results and phenotypic susceptibility testing, we found an overall concordance of 90.2% with the line probe data. Three isolates among 51 RIF-resistant isolates examined had no mutation in the 81-bp rpoB hotspot and two had insertional mutations that cannot be identified by the line probe kit [64]. Higher extents of correlation between the line probe kit results and RIF resistance have been reported elsewhere [6]. Mutations in rpoB have been evaluated using another commercial kit, the MutationScreener Kit (Ambion, Inc., Austin, Texas), which operates on the

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principle of nonisotopic RNase cleavage assay [65]. Target DNA is amplified by PCR using primers with 20-base phage promoters on their 59-ends followed by transcription of the products by phage RNA polymerase and hybridization to form double-stranded RNA. Mutations in rpoB result in basepair mismatches when complementary reference (wild type) and mutant transcripts are hybridized. After hybridization, the duplex RNA targets are treated with a mixture of RNases capable of cleaving basepair mismatches on both strands. The double-stranded cleavage products are separated on a simple native agarose gel and detected by ethidium bromide staining under UV light. In an evaluation of 46 isolates in one research laboratory, sensitivity and specificity were found to be 100% and 96%, respectively [65]. One of the most studied noncommercial methods that has been evaluated in research laboratories for identifying mutations associated with antituberculosis drug resistance is single-strand conformation polymorphism (SSCP) electrophoresis [66]. Any two PCR products that are 400 bp or longer and have at least one basepair difference in their sequences will have different electrophoretic mobilities in vertical nondenaturing acrylamide gels if the appropriate conditions are met. These conditions include gel strength and temperature, voltage and run length, and how the PCR products are pretreated. The products are typically heated to 88 to 95 C for 4 min in the presence of a strong denaturant (eg, methyl mercury hydroxide or formamide) and immediately cooled on ice before loading the gel. Samples labeled with a radioisotope (32P) were first used to screen for rpoB mutations by Telenti in 1993 [67]. Nonradioactive versions of the SSCP method have since been described for screening resistance mutations for other antituberculosis drugs [51,54,68–71], and it has been shown to be capable of identifying resistant subpopulations of rpoB mutants [68]. Although the SSCP method has been also performed using a high throughput capillary sequencer [68], it is most often performed with relatively inexpensive electrophoresis apparatuses and reagents. Because mobility shifts for some mutations are minor, it is important to space wild-type controls for comparison at regular intervals in gels, and suspect isolates must be tested for resistance by a more established method. Although the SSCP method has the potential to detect novel mutations within PCR products, it is possible that some mutations will cause imperceptible or negligible shifts. For larger genes having diverse mutations, such as pncA, SSCP may be unsuitable. Other genotypic methods that offer the potential to identify novel as well as previously documented mutations are denaturing gradient gel electrophoresis (DGGE), temperature-mediated heteroduplex analysis (TMHA) using DNA high performance liquid chromatography (HPLC), and branch migration inhibition (BMI) [72–74], all of which require annealing of target and reference DNA strands. The DGGE method was shown to identify at least 14 rpoB mutations, results for 117 isolates were completely concordant with DNA sequence analysis and

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RIF susceptibility testing, and patterns could be generated using DNA directly extracted from culture-positive processed sputum samples [74]. The TMHA/DNA-HPLC method was shown to be capable of identifying 19 mutations associated with resistance to five primary antituberculosis drugs as well a polymorphism on oxyR used to differentiate some MTC species [72,75]. However, instrumentation for these methods is expensive, and standardization of both procedures in individual laboratories may pose some difficulties. Stable, four-stranded cruciform structures are formed using the BMI method if a mutation is present in a DNA strand relative to a reference strand, both of which are amplified using specially-designed primers. The cruciform structures are undetectable (eg, when using chemiluminescent labels) if test and reference strands are homologous. Liu et al [73] evaluated 83 M tuberculosis isolates for RIF and PZA resistance using BMI and obtained complete agreement with phenotypic susceptibility results. Other technologies that have been shown to be useful for identifying antituberculosis drug resistance markers are oriented toward identifying specific nucleotide polymorphisms. Technologies that conform to this principle and that have been applied to genotypic antituberculosis drug testing include real-time fluorescent PCR in the LightCycler instrument [76,77], structure specific cleavage [78], and molecular beacons [79,80]. The latter of these assays employs multiple complex fluorogenic compounds bound to specific oligonucleotides, all of which are quenched if probes are in free solution (unhybridized). Using target DNA extracted directly from clinical specimens, as many as five regions may be simultaneously interrogated within a single tube in less than 3 hours. Piatek et al [80] demonstrated the practical application of molecular beacons to genotypic antituberculosis drug susceptibility testing of 149 M tuberculosis isolates by performing real-time multiplex PCR in closed tubes using four molecular beacons for mutations in rpoB for RIF resistance and in katG, inhA, and ahpC-oxyR for INH resistance. Sensitivity and specificity for INH resistance were 85% and 100%, respectively; they were 98% and 100%, respectively, for RIF resistance [80]. High-density oligonucleotide arrays (DNA microarrays) is a recently introduced technology that has already proven useful for whole genome analyses of mycobacteria, particularly for gene expression experiments [81,82] and in locating specific repetitive elements such as IS6110 [83]. Microarrays also show promise for genotypic antituberculosis drug susceptibility testing [60]. Another technology that employs microchips was recently used to identify mutations in rpoB among 30 RIF-resistant M tuberculosis isolates [84]. The mutations were identified using hybridization to a set of 42 oligonucleotides bound to a microchip as well as by allele-specific on-chip PCR and on-chip ligase detection reactions. The ligase detection reaction assay was also capable of identifying mutations in mixtures containing 1% resistant and 99% susceptible bacilli [84].

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Molecular typing methods for M tuberculosis Before the late 1980s, laboratory support for epidemic investigations involving M tuberculosis most often involved only antimicrobial susceptibility profiles and, perhaps, bacteriophage types, both of which were obtained using very time-consuming and laborious procedures and were often of minimal use in the investigation of outbreaks [85]. The use of repetitive genetic elements as probes to determine strain-specific heterogeneities in the M tuberculosis genome was introduced in 1988 by Eisenach et al [86] and quickly became a standard epidemiologic typing tool that has contributed greatly in TB control efforts throughout the world [85]. The repetitive element, IS6110 [87], is present in 0 to 30 copies in the genome of MTC strains, although single copy strains often are identified as M bovis. The IS6110 typing method was standardized in 1993 [88] and involves digesting DNA isolated from cultures with PvuII. These fragments are electrophoresed, blotted to hybridization membranes, and probed with labeled IS6110 PCR products to reveal restriction fragment length polymorphism (RFLP) patterns or fingerprints that can be analyzed to determine the relationships among isolates under epidemiologic investigation. Banding patterns for isolates that may be epidemiologically linked are often subjected to cluster analysis (eg, using software that generates a dendrogram that shows relatedness of the patterns). Isolates that have all bands matching may be considered clonal or the same strain, suggesting possible transmission linkages. An amplification-based version of IS6110 typing (mixed linker PCR) was introduced in 1993 by Haas et al [89] and offers a level of strain resolution comparable with IS6110 RFLP typing. The protocol for this procedure requires multiple steps including restriction with HhaI, ligation of uracilcontaining oligonucleotide linkers, treatment with uracil DNA glycosylase, nested PCR, and finally electrophoretic separation of DNA bands. A method that requires fewer procedural steps is spacer oligonucleotide typing (spoligotyping), which was introduced in 1997 [90]. This method evaluates the presence of 43 unique spacer sequences that are each flanked by identical 36-bp direct repeats in the MTC genome. The region is amplified by PCR using primers to the direct repeats and hybridized to the 43 spacer oligonucleotides that have been immobilized on a hybridization membrane. One of the spoligotyping primers is labeled with biotin and identification of hybrids is performed by a chemiluminescent reaction using a streptavidineperoxidase conjugate. The 43 hybridization results are then converted to a 15-digit octal numeric code that denotes the strain type [91]. Because spoligotyping is an amplification-based method, considerably less target DNA is required than for IS6110 RFLP typing. The use of hybridization membranes for spoligotyping may be avoided by using high throughput flow cytometers such as those produced by Luminex Corporation (Austin, Texas), which offer multiplex hybridizations on polystyrene microspheres,

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each of which may be differentiated by the instrument. Each of the 43 spacer oligonucleotide probes may be attached to a microsphere available from a set of 100 microspheres, each with a different color spectrum, and hybrids formed with the spacer region amplicons may be detected and quantitated automatically. In general, spoligotyping will not resolve collections of M tuberculosis into more clusters than IS6110 RFLP typing except for those isolates with fewer than five copies of the insertion element [92,93]. Another PCR-based typing method for M tuberculosis isolates is mycobacterial interspersed repetitive unit (MIRU) typing, which is a variable number tandem repeat method [94]. The approach exploits 12 regions of the M tuberculosis genome, each of which has a number of tandem nucleotide repeats ranging in size from 53 to 77 bp and from 1 to 11 copies in each region. Twelve sets of PCR primers are required, whereas only a single set of primers is needed for spoligotyping. No hybridizations are required, however, for MIRU typing; the size (basepairs) of each PCR product is used to determine the number of repeats at each locus, which is then used to generate a 12-digit strain type identifier. Whether MIRU typing offers powers of strain discrimination comparable to that of IS6110 RFLP typing has not yet been established, except for isolates having low copy numbers of IS6110. Cowan et al [95] evaluated 180 M tuberculosis isolates, each with fewer than seven copies of IS6110, and found more distinct MIRU patterns (80 patterns) than IS6110 RFLP patterns (58 patterns). Because sizes of PCR products are used to determine MIRU typing patterns, instruments that automate electrophoresis and size determinations of dsDNA may be useful. Multiplex MIRU typing using fluorescently labeled primers and an automatic sequencer has been described [96], and the use of high-throughput, fluoresence-based capillary instruments such as the Beckman CEQ8000 (Beckman Coulter, Inc., Fullerton, California) or the ABI3100 (Applied Biosystems, Inc., Foster City, California) for MIRU typing is feasible. Identification of nontuberculous mycobacteria Nontuberculous mycobacteria (NTM), also referred to as mycobacteria other than M tuberculosis, may also be seen using AFB smear microscopy and may be grown on either egg-based or agar-based media or in brothbased systems. Because NTM organisms may become overgrown with contaminants, environmental samples (eg, water or soil) and some clinical specimens (eg, sputum or stool) must be decontaminated before culture, despite the probability that the numbers of desired viable mycobacterial organisms will be reduced as a result, a problem that is also encountered with tubercle bacilli [1]. Members of some species, however, require growth supplements or culture temperatures other than 35 C to 37 C [1]. In addition to optimal growth temperature, the identification of nontuberculous members of this genus has been built upon parameters including staining

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properties, colony morphology, photochromogenicity, ability to grow upon various selective media, animal pathogenicity, and biochemical tests including citrate, inositol, mannitol, and sorbitol use, nitrate reduction, iron uptake, and aryl-sulfatase activity [1,5]. Analyses of mycolic acid patterns using HPLC was proposed as an alternative means to identify mycobacteria in 1991 [97] and has since become a standard identification tool [98]. Extraction of the mycolic acids from culture is a straightforward heat saponification, acidification, and methanolic extraction process followed by derivitization of the fatty acids with UV-absorbing esters. Detection and characterization of mycolic acids extracted directly from clinical specimens also is possible through the use of fluorescent labeling compounds [99]. Genotypic identification of mycobacteria was introduced at a commercial level by GenProbe, Inc., San Diego, California as probe kits (now available as the Accuprobe System) for the identification of M avium, M avium complex (MAC), M intracellulare, M gordonae, and M kansasii, as well as MTC organisms. These assays use specific DNA probes that are labeled with an acridinium ester detector molecule that emits a chemiluminescent signal and are used in a hybridization protection assay to target rRNA. The Accuprobe kit has been reported to be both sensitive and specific [62]—notably by LeBrun et al [100], who evaluated 134 clinical mycobacterial isolates including 36 MTC, 40 M avium complex, 27 M gordonae, 9 M kansasii, and 22 unidentified Mycobacterium spp isolates. Specificity was found to be 100%, and sensitivity was 95.2% or greater for four probes that were evaluated [100]. A noncommercial genotypic scheme to identify a larger assortment of Mycobacterium species was described in 1993 [101] and has since been designated hsp65 PCR restriction analysis (PRA). The assay is based on polymorphisms in the gene encoding the 65-kd heat shock protein [102], which appear to be stable features of Mycobacterium species [103]. The polymorphisms are most often identified by digesting PCR products of hsp65 with BstEII and HaeIII and determining sizes of the fragments. The method is considered a research tool and has not been cleared by the United States FDA for use in clinical laboratories. The hsp65 PRA method will not distinguish MTC species from one another or some NTMs (eg, M avium from M avium subspecies M paratuberculosis). Some hsp65 PRA patterns show only minor differences in fragment sizes for some species, which strongly suggests the need for more precise fragment sizing than is typically available from estimating sizes by comparison with size standards in electrophoretic gels. A multicenter collaboration has established a website (http://www.hos pvd.ch/prasite) with a database for mycobacterial hsp65 restriction patterns, including an algorithm into which experimental restriction data can be entered and analyzed for identification of Mycobacterium species. Restriction analysis of a variable portion of the 16S rRNA gene has also been employed as a research tool to identify Mycobacterium species.

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Avaniss-Aghajani et al [104] amplifed 475-bp PCR products from this gene region using a fluorescent-labeled primer for the sense strand, restricted the products with five enzymes, and used an automated slab gel sequencer to show that 13 Mycobacterium species could be identified. A major advantage of this identification method was the development of software to automatically determine fragment sizes that reduced operator errors in measurements of fragment sizes. Hernandez et al [105] combined aspects of hsp65 and 16s rRNA PRA to develop a scheme in which 19 Mycobacterium species were identified using a fluorescent capillary sequencer (ABI model 310). The hsp65 and 16S rRNA genes were amplified using primers labeled with a different fluorescent dye for each gene. Each product was digested with two restriction enzymes and, because only the end-most fragments were labeled, 8 or fewer fragments could be identified [105]. Restriction analysis of hsp65 with enzymes other than HaeIII and BstEII and analysis of the 16S-23S rRNA spacer region have also been shown to be useful tools for Mycobacterium species identification [106–108]. Two commercial kits to identify mycobacteria based on reverse hybridization of ribosomal DNA regions have been recently introduced. The GenoType assay (Hain Lifescience, Nehren, Germany) targets 23S rDNA and the INNO-LiPA Mycobacteria kit (Innogenetics, Ghent, Belgium) targets the 16S-23S spacer region. Both kits have probes to identify MTC, M avium, M intracellulare, M kansasii, M chelonae, M gordonae, M xenopi, and M scrofulaceum organisms. The LiPA kit also differentiates two members of the M avium complex (M avium and M intracellulare) and some subspecies of M chelonae and M kansasii, while the GenoType kit can also identify M malmoense, M celatum, M peregrinum, M phlei and two subspecies of M fortuitum. A recent comparison of the two kits showed that 89.4% and 95.1% of 81 mycobacterial isolates were correctly identified by the LiPA and the GenoType kits, respectively [109], while another study on a larger collection (238 strains, including 143 isolates representing the targeted groups or species and 95 isolates representing 25 additional Mycobacterium groups or species) demonstrated 99.6% accuracy with the LiPA kit [110]. Sequence analysis of the signature region (500 bp) of the 16S rRNA gene is often used for mycobacterial taxonomy [111]. In some instances, a larger 16S rDNA region (1540 bp) must be sequenced to resolve other conflicting data (eg, HPLC patterns) to identify unusual isolates.

Molecular typing of nontuberculous mycobacteria Infections caused by NTM species have been reported with increasing frequency since the mid-1990s [1,2], underscoring the need for improved methods for identification and strain characterization. Classic typing methods such as phage typing, antibiograms, biochemical profiling, and serotyping have yielded to molecular methods. Plasmid profiling has little

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application for typing most NTMs, and insertion element polymorphic typing similar to IS6110 RFLP fingerprinting of M tuberculosis has proven to be of limited use, except for IS1245 typing of M avium [112]. The most frequently used typing methods for most NTM species are multilocus enzyme typing (MET) [113], pulse-field gel electrophoresis (PFGE) of infrequent-target restriction enzyme digests [114], and randomly-amplified polymorphic DNA (RAPD) electrophoresis [115]. The MET method is performed by starch gel electrophoresis of extracts (most often impregnated into small pieces of filter paper) of organisms containing housekeeping enzymes such as esterases and dehydrogenases. The gels are sliced in a horizontal plane, and the slices are placed into solutions containing stains capable of identifying each enzyme. Isoenzymes of these proteins often have strain-specific electrophoretic mobilities that are useful for differentiating isolates [116]. Large restriction fragment examination by PFGE requires extraction of DNA in a fashion that minimizes nonspecific shearing followed by digestion with restriction enzymes with few recognition sites in mycobacterial genomes, such as SmaI, XbaI, or DraI [114]. These digestions are most often performed in small blocks of solidified agarose to stabilize the DNA even further before agarose electrophoresis in apparatuses such as the Chef-DR III horizontal system (BioRad Laboratories, Richmond, California), which may be programmed to alternate the electrophoretic fields around the agarose gels that facilitate the movement of large (48 kb) dsDNA fragments. The RAPD method is perhaps the easiest typing method for NTM species, but reproducibility of electrophoretic patterns has not been established. Crude DNA preparations are used as templates in low stringency PCR in which the annealing temperature is considerably lowered (eg, to 37 C) and only a single short (10-mer) primer is used. Banding patterns of the samples are compared after electrophoresis; as for PFGE, DNA is stained using ethidium bromide. The most reliable data are obtained if all isolates being evaluated are amplified and electrophoresed together in the same run and strengthened even more by using multiple runs, each with a different primer.

Summary Just as tuberculosis has persisted for many centuries as one of most serious and deadly infectious diseases in many parts of the world, so has the motivation to develop improved laboratory methods for characterizing M tuberculosis isolates. Modern technology has lead to great improvements in mycobacteriology laboratory procedures, particularly in detection, identification, epidemiologic strain typing, and drug susceptibility testing. Although the usefulness of some of these newer methods is under evaluation, many already are showing potential as adjuncts to clinical diagnostic procedures.

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