The Mycobacterium tuberculosis complex (MTC) comprises closely related species responsible for strictly human and zoonotic tuberculosis. Accurate species determination is useful for the identification of outbreaks and epidemiological links. Mycobacterium africanum and Mycobacterium canettii are typically restricted to Africa and M. bovis is a re-emerging pathogen. Identification of these species is difficult and expensive.
The Exact Tandem Repeat D (ETR-D; alias Mycobacterial Interspersed Repetitive Unit 4) was sequenced in MTC species type strains and 110 clinical isolates, in parallel to reference polyphasic identification based on phenotype profiling and sequencing of pncA, oxyR, hsp65, gyrB genes and the major polymorphism tandem repeat. Inclusion of M. tuberculosis isolates in the expanding, antibiotic-resistant Beijing clone was determined by Rv0927c gene sequencing. The ETR-D (780-bp) sequence unambiguously identified MTC species type strain except M. pinnipedii and M. microti thanks to six single nucleotide polymorphisms, variable numbers (1–7 copies) of the tandem repeat and two deletions/insertions. The ETR-D sequencing agreed with phenotypic identification in 107/110 clinical isolates and with reference polyphasic molecular identification in all isolates, comprising 98 M. tuberculosis, 5 M. bovis BCG type, 5 M. canettii, and 2 M. africanum. For M. tuberculosis isolates, the ETR-D sequence was not significantly associated with the Beijing clone.
ETR-D sequencing allowed accurate, single-step identification of the MTC at the species level. It circumvented the current expensive, time-consuming polyphasic approach. It could be used to depict epidemiology of zoonotic and human tuberculosis, especially in African countries where several MTC species are emerging.
The Mycobacterium tuberculosis complex (MTC) comprises several closely related species responsible for strictly human and zoonotic tuberculosis. Some of the species are restricted to Africa and were responsible for the high prevalence of tuberculosis. However, their identification at species level is difficult and expansive. Accurate species identification of all members is warranted in order to distinguish between strict human and zoonotic tuberculosis, to trace source exposure during epidemiological studies, and for the appropriate treatment of patients. In this paper, the Exact Tandem Repeat D (ETR-D) intergenic region was investigated in order to distinguish MTC species. The ETR-D sequencing unambiguously identified MTC species type strain except M. pinnipedii and M. microti, and the results agreed with phenotypic and molecular identification. This finding offers a new tool for the rapid and accurate identification of MTC species in a single sequencing reaction, replacing the current time-consuming polyphasic approach. Its use could assist public health interventions and aid in the control of zoonotic transmission in African countries, and could be of particular interest with the current emergence of multidrug-resistant and extended-resistance isolates.
Citation: Djelouadji Z, Raoult D, Daffé M, Drancourt M (2008) A Single-Step Sequencing Method for the Identification of Mycobacterium tuberculosis Complex Species. PLoS Negl Trop Dis 2(6): e253. https://doi.org/10.1371/journal.pntd.0000253
Editor: Pamela L. C. Small, University of Tennessee, United States of America
Received: October 11, 2007; Accepted: May 20, 2008; Published: June 18, 2008
Copyright: © 2008 Djelouadji et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by Unité des Rickettsies - CNRS UMR 6236 UMR_D 198 and l'Oeuvre Antituberculeuse des Bouches-du-Rhône, Marseille, France. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors declare that they are the inventors of a protective patent deposited by the Mediterranée University, Marseilles, France on this matter.
The Mycobacterium tuberculosis complex (MTC) comprises several closely related species responsible for strictly human and zoonotic tuberculosis (Figure 1). In addition to M. tuberculosis, which represents the leading cause of human tuberculosis worldwide and is now emerging as extensively drug-resistant tuberculosis strains , other MTC species have been found in patients, typically in African countries (Figure 2). Mycobacterium bovis is a re-emerging, zoonotic agent of bovine tuberculosis  whose prevalence probably depends on variations in direct exposure to cattle and consumption of unpasteurised dairy products . The prevalence of Mycobacterium africanum type I (West Africa) and type II (East Africa)  has decreased in several African countries over the last decades ,. Mycobacterium canettii, a rare MTC species, has been isolated recently in patients exposed in Africa . Mycobacterium microti, a vole and small rodent pathogen  that is closely related to the so-called Dassie-bacillus and infects small mammals in South Africa and the Middle East ,, has been isolated in humans . Mycobacterium caprae is a rare cause of tuberculosis in cattle , and zoonotic tuberculosis in humans  while Mycobacterium pinnipedii has been isolated from seal lions and fur seals . A recent description of the re-emergence of M. bovis in cattle, along with the direct interhuman transmission of this zoonotic organism  in a six-case cluster that included one death in United Kingdom , illustrates the potential of emerging and re-emerging zoonotic tuberculosis due to MTC species other than M. tuberculosis and the necessity for accurate species identification.
Green circles, M. canettii; blue circles, M. caprae; pink circles, M. microti; orange circles, M. pinnipedii.
Based on a review of papers published in 1999–2007 , –. For each country, prevalence rates and reference are indicated in cartoons. Mtub, M. tuberculosis; M afr, M. africanum; Mbov, M. bovis; Mcan, M. canettii; MTC, Mycobacterium tuberculosis complex.
Accurate species identification of all MTC members is warranted in order to distinguish between strict human and zoonotic tuberculosis and to trace source exposure during epidemiological studies. Indeed, phenotypic methods of identification relying on colony morphology, oxygen preference, niacin accumulation, nitrate reductase activity, growth kinetics and resistance to thiophene-2-carboxylic acid hydrazide (TCH) and PZA  are hampered by slow growth of MTC members and subjective interpretation of colony morphology and cross-resistance to drugs . They do not always allow unambiguous species identification in every case. Recent studies of MTC species responsible for animal and human tuberculosis in tropical countries have relied on molecular methods including mycobacterial interspersed repetitive-unit-variable-number tandem-repeat (MIRU-VNTR) typing, IS6110-RFLP and spoligotyping –. Molecular differentiation of MTC members has been complicated by low sequence variability at the nucleotide level, illustrated by a 85–100% DNA/DNA relatedness and a 99–100% 16S rDNA sequence similarity ,. Nucleic acid-based assays such as acridinium ester-labelled DNA probes (AccuProbe; Gene Probe Inc, San Diego, CA) have proven to be reliable tools for assigning an isolate to the MTC ,, but they do not allow for identification at the species level. Molecular identification based on deleted regions (RD), RD1, RD9 and RD10 , are limited by the necessity of interpreting negative results in the case of the absence of a specific deletion. The detection of single nucleotide polymorphisms (SNP) in the pncA gene , the oxyR locus , the mtp40 gene , and the restriction fragment length polymorphism of the hupB gene  differentiated M. tuberculosis from M. bovis but not from other MTC species. The major polymorphism of tandem repeat (MPTR) sequencing differentiated M. tuberculosis (Sequevar long), M. bovis and M. microti (Sequevar Med-G), M. bovis BCG (Sequevar Med-C) and M. africanum (Sequevar short), but other MTC species were not studied . The gyrB gene proved to be an effective target ,, as an identification scheme has been proposed based on Pyrosequencing analysis of four single nucleotide polymorphism (SNPs) in gyrB , and a DNA strip based on gyrB is commercially available (HAIN Genotype MTBC DNAstrip test, Hain Lifescience, Nehren, Germany) . Both approaches, however, fail to differentiate M. tuberculosis from M. africanum type II and M. canettii; and M. africanum type I from M. pinnipedii. IS6110-RFLP, VNTR typing and Spoligotyping , emerged as reference methods to study the diversity of MTC species in resource-limited countries, despite the fact that these methods may not recognize rarely encountered species and may not appreciate the entire genetic diversity of strains, as they are not based upon the sequencing of molecular targets .
When investigating intergenic spacers in the genotyping of M. tuberculosis, we found that one spacer, previously identified as the Exact Tandem Repeat D (ETR-D)  and aliased Mycobacterial Interspersed Repeat Unit 04 (MIRU04) , exhibited a variable sequence among M. tuberculosis isolates. Analysis of this spacer had been previously shown to distinguish between M. bovis and the M. bovis BCG type . We therefore further investigated whether sequencing the ETR-D could identify all of the MTC at the species level. In this study, we demonstrate that ETR-D sequencing offers a new tool for the rapid and accurate identification of MTC species in a single reaction.
M. tuberculosis CIP103471, M. bovis CIP105050, M. africanum CIP105147T (type I), M. bovis BCG vaccine strain type 105060, M. microti CIP104256T, M. canettii CIP140060001T, M. pinnipedii ATCC BAA-688, and M. caprae CIP105776T reference strains were purchased from the Collection Institut Pasteur (CIP, Paris, France) and American Type Culture Collection (ATCC, Rockville, USA). The following non-tuberculosis mycobacteria were tested in order to assess the specificity of ETR-D spacer sequencing: Mycobacterium avium IWGMT49 T, Mycobacterium intracellulare CIP104243 T, Mycobacterium chimaera CIP107892 T, Mycobacterium colombiense CIP108962 T, Mycobacterium haemophilum CIP105049 T, Mycobacterium ulcerans CIP105425 T, Mycobacterium xenopi CIP104035 T, Mycobacterium abscessus CIP104536 T, Mycobacterium chelonae CIP104535 T, Mycobacterium fortuitum ATCC49404 and Mycobacterium mucogenicum CIP 105223 T. Quality of DNA was controlled by parallel partial rpoB PCR amplification as previously described . One hundred and ten MTC clinical isolates (Table 1) recovered from Microbiology Laboratory in Marseille (n = 102), from Institut Pasteur in Djibouti (n = 3) and from Institut de Pharmacologie et Biologie Structurale, Toulouse (n = 5) were also analyzed. All isolates were identified as members of the MTC by phenotypic characterization and a gene probe assay according to the manufacturer (AccuProbe; Gene Probe Inc, San Diego, Calif). This study was approved by the ethics committee of the Institut Féfératif de Recherche 48, Marseilles, France.
Phenotypic characterisation included colony morphology, a urease test controlled after 3 and 18 hour incubation, and oxygen consumption measured after inoculation of a 0.2 ml actively growing mycobacterial suspension into 40 ml of Middlebrook 7H10 into the Bactec 9000MB system (Becton and Dickinson, Le Pont de la Claix, France) after a 3-week incubation. Drug susceptibility tests for thiophene-2-carboxylic acid hydrazide (TCH) and PZA were performed as previously described .
Reference tests for molecular identification
The identification of reference strains and clinical isolates identified as M. bovis BCG type, M. canettii and M. africanum by ETR-D sequencing (see below) was confirmed by parallel reference molecular tests. Every isolate coated on beads was inactivated as previously described  and the DNA was extracted using a Qiagen kit (Qiagen, Courtaboeuf, France). DNA was used as a template for PCR amplification of pncA, oxyR, hsp65, gyrB genes and sequence analysis of MPTR was performed as previously described , , –, In addition, we sequenced the Rv0927c-pstS3 intergenic region in all clinical isolates identified as M. tuberculosis in order to identify the Beijing genotype . Amplified products were visualized by agarose gel electrophoresis and direct sequencing was performed as described above. Sequences were edited using the Auto assembler program (Applied Biosystems, Courtaboeuf, France) and aligned using CLUSTAL W (http://pbil.ibcp.fr). Original sequences were deposited into GenBank (http://www.ncbi.nlm.nih.gov/sites/entrez/).
ETR-D spacer sequencing
Amplification and sequencing of the ETR-D spacer located between the putative histidine kinase Senx3 upstream and the sensory transduction protein Regx3 downstream were done using direct primers: 5′-GTTGATCGAGGCCTATCACG-3′ and 5′-GAATAGGGCTTGGTCACGTA-3′. The PCR mixture contained 33 µl H2O, 5 µl 10× buffer (Qiagen), 2 µl 25× MgCl2, 5 µl 10× dDNTP, 1 µl forward primer, 1 µl reverse primer, 0.25 µl hotstart Taq (Qiagen) and 2 µl target DNA. Appropriate negative controls consisting of PCR mix without target DNA were also included. PCRs were performed using the following program: 15 min enzyme activation at 95°C, followed by 34 cycles consisting of 95°C for 30 s, 58°C for 30 s, 72°C for 1 min, followed by a 5 min elongation step at 72°C. After agarose gel electrophoresis, PCR products were purified and subjected to sequencing in both directions by using the BigDye Terminator 1.1 Cycle Sequencing kit (Applied Biosystems). Sequencing electrophoresis was performed on a 3130 genetic analyzer (Applied Biosystems). The sequences were edited using the Auto assembler program (Applied Biosystems) and aligned using CLUSTAL W (http://pbil.ibcp.fr). Original ETR-D sequences were deposited into Genbank (http://www.ncbi.nlm.nih.gov/sites/entrez/).
Phenotypic identification (Table 2)
As for reference strains, M. tuberculosis exhibited eugonic growth that was inhibited by the presence of PZA but not by TCH and showed aerophilic growth on Middlebrook agar positive urease at 18 hours. M. bovis and M. bovis BCG type strains exhibited microaerophilic dysgonic growth, did not grow in the presence of TCH, but were resistant to PZA and exhibited a positive urease activity at 3 hours for M. bovis BCG type and at 18 hours for M. bovis. M. africanum type I differed from M. bovis by its susceptibility to PZA. M. canettii exhibited eugonic growth in the presence of PZA and TCH, and showed a positive urease activity at 3 hours and aerophilic growth on Middlebrook. M. microti, M. capare and M. pinnipedii exhibited eugonic growth that was inhibited by TCH and PZA, and a positive urease at 18 hours. As for clinical isolates, 101/110 of isolates were phenotypically identified as M. tuberculosis, 5 as M. canettii, 3 as M. bovis BCG type and one as M. africanum.
Reference molecular identification (Table 3)
In all PCR experiments, negative controls remained negative. All reference strains and clinical isolates yielded an amplicon of the expected size when amplified for pncA, oxyR, hsp65, gyrB genes, Rv0927c-pstS3 intergenic region and MPTR. By comparison with M. tuberculosis, the 410-bp oxyR gene sequence exhibited a previously known A285G polymorphism in M. bovis and M. bovis BCG type  and a newly identified T136G polymorphism in M. canettii. The 561-bp pncA gene sequence exhibited a previously known G253C polymorphism in M. bovis and M. bovis BCG type  and a G222A polymorphism in M. canettii . The 441-bp hsp65 gene exhibited a previously known T235C polymorphism in M. canettii  and a newly identified G376C polymorphism in M. africanum type I. The 1.020-bp gyrB gene sequence exhibited an identical sequence in M. tuberculosis, M. canettii and M. caprae, a previously known A756G polymorphism in M. bovis and M. bovis BCG type, a T675C polymorphism in M. microti, and an identical sequence was identified in common with M. africanum type I and M. pinnipedii . Sequence analysis of MPTR (300-bp) exhibited a unique sequence for M. tuberculosis (Sequevar Long), M. africanum type I strain (Sequevar Short), M. bovis and M. microti (Sequevar MED-G) and M. bovis BCG type (Sequevar MED-C) ; MPTR sequencing newly identified C99T, G164C and A267G polymorphisms in M. canettii; the M. caprae strain exhibited Sequevar Long in common with M. pinnipedii and M. tuberculosis reference strains. Original sequences found in this study were deposited in GenBank under the following accession numbers (GenBank: EF656461, EF 656463, EF 656464).
Sequence analysis of clinical isolates using the five previous targets yielded four different profiles. One profile comprised 98 isolates identified as M. tuberculosis, including three isolates identified as W-Beijing strains using a G127A polymorphism in Rv0927c-pstS3 intergenic region, a second profile comprised five isolates identified as M. bovis BCG type; a third profile included five isolates identified as M. canettii and a fourth profile included two isolates identified as M. africanum type I.
ETR-D sequence analysis
For all ETR-D experiments, negative controls remained negative. All the non-tuberculosis mycobacteria yielded a negative ETR-D PCR amplification whereas they yielded an amplicon of the expected size through rpoB PCR amplification. The size of PCR products obtained with MTC reference strains varied from 497-bp for M. canettii, 545-bp for M. bovis, 564-bp for M. caprae, 598-bp for M. bovis BCG type, 651-bp for M. africanum type I, 805-bp for M. tuberculosis and 959-bp for M. microti and M. pinnipedii. Each of the eight reference strains exhibited a unique ETR-D sequence, exhibiting one to seven copies of a tandem repeat, six mutations and two deletions/insertions.
M. tuberculosis exhibited three different alleles combining two or five copies of a 77-bp repeat unit and one T/G SNP at the fifth base of the tandem repeat, in addition to one 53-bp repeat unit. The M. tuberculosis reference strain exhibited five copies of a 77-bp repeat unit followed by one 53-bp repeat unit copy; M. microti and M. pinnipedii exhibited seven copies of a 77-bp repeat unit and one 53-bp repeat unit, M. africanum type I exhibited three copies of a 77-bp repeat unit and one 53-bp repeat unit, in addition to one T/C polymorphism at position 75. M. bovis exhibited four copies of a 77-bp repeat unit and one 53-bp repeat unit in addition to an A/G SNP at position 773; M. bovis BCG type exhibited three copies of a 77-bp repeat unit in addition to an A/G SNP at position 773. M. caprae exhibited three copies of a 77-bp repeat unit, a 34-bp deletion and one 53-bp repeat unit. M. canettii exhibited one copy of a 77-bp repeat unit and three SNPs following the tandem repeat in addition to a 53-bp repeat unit. ETR-D sequences of MTC type strains were deposited in GenBank under the following accession numbers (GenBank: EU180228-EU180234). ETR-D sequencing identified 98/110 MTC clinical isolates as M. tuberculosis including 45 isolates presenting allele 1, 26 isolates presenting allele 2, and 27 isolates presenting allele 3; the ETR-D allele did not correlate with Beijing genotype (P = 0.2). 5/110 isolates were identified as M. bovis BCG type, 5/110 isolates as M. canettii, and 2/110 isolates as M. africanum type I. All unique ETR-D sequences were deposited into our freely available database at http://ifr48.timone.univ-mrs.fr/MST_Mtuberculosis/mst.
Comparison between ETR-D identification and reference phenotypic and molecular identifications of clinical isolates
ETR-D identification was in agreement with phenotypic identification in 107/110 (97.27%) of clinical isolates. Three isolates phenotypically identified as M. tuberculosis were identified by ETR-D sequencing and reference molecular methods as M. bovis BCG type in two cases and M. africanum type I in one case. Reference molecular identification agreed with ETR-D identification in 100% (110/110) of clinical isolates.
Previous methods for MTC species identification either combined the amplification of several genomic regions in order to identify all species , or analyzed one gene polymorphism to distinguish between only two species. ETR-D spacer sequencing herein developed proved to be specific for the MTC and allowed the differentiation of the 7/8 MTC species in a single reaction. Indeed all the non-tuberculosis mycobacteria yielded a negative ETR-D PCR amplification as previously described .
The fact that M. africanum type II was not included in the present study may not modify this conclusion. In fact, the taxonomic status of M. africanum type II has been disputed , but it is now regarded as a phenotypic variant of M. tuberculosis (genotype Uganda) ,. ETR-D sequencing agreed in all cases with reference molecular identification. In this study, new mutations were identified because some genes were sequenced for the first time in some MTC species including the oxyR gene and MPTR in M. canettii and the hsp65 gene in M. africanum type I (Table 3). ETR-D sequencing revealed that 3/110 clinical isolates identified as M. tuberculosis by phenotypic tests comprised two M. bovis BCG type isolate and one M. africanum type I isolate. The 497-959-bp size of ETR-D allows one-step sequencing using a modern capillary sequencer and software and may be easily sequenced using Pyrosequencing and additional internal primers. Cost was decreased in comparison with the current polyphasic approach and any microbiologist could compare the ETR-D sequence with those that we deposited in the versatile, freely accessible databank at http://ifr48.timone.univ-mrs.fr/MST_MTuberculosis/mst. This identification technique, based on PCR amplification, could be directly applied to clinical specimens exhibiting acid-fast bacilli.
ETR-D sequence identification relied not only on the variation in the number of tandem repeats illustrated by various PCR product sizes, as previously described  for M. tuberculosis, M. africanum, M. bovis group , but also on specific SNPs, which are stable events  accounting for 55.5% of genetic events observed in this study and on insertion/deletion events (accounting for 22.2% of genetic events). However, the ETR-D sequence was not correlated with the Beijing genotype as defined by Rv0927c-pstS3 intergenic region sequencing. This indicates that, although 3 ETR-D genotypes were found among M. tuberculosis isolates in this study, ETR-D sequencing alone cannot be used for genotyping. It is not surprising that the same, limited genomic region does not have the potential to identify at the species and strain levels. ETR-D sequencing provides, for the first time, a unique sequencing test capable of distinguishing all MTC species in a single step.
Accurate identification of MTC isolates at the species level is of particular interest in Africa where species other than M. tuberculosis were characterized in human tuberculosis and M. bovis remains a huge problem for cattle  (Figure 2). Their identification may direct specific epidemiological investigation. In Africa, the prevalence of M. bovis in human tuberculosis was correlated with the prevalence in the local cattle population . Consumption of unpasteurised milk and of poorly heat-treated meat, and close contact with infected animals represent the main sources of infection for humans . However, human to human transmission of M. bovis was recently reported in a 6-case cluster including one death due to M. bovis meningitis in United-Kingdom . In addition, the emergence of MDR M. bovis has been documented, raising infection control in health care settings ,. M. bovis BCG type derived from the closely related virulent M. bovis after 230 serial passages had led to a considerably increased rate of disseminated BCG disease in HIV-infected infants reported in South Africa , although diagnoses were based on a few biochemical tests including the urease test and RD1deletion . ETR-D sequencing allows unambiguous distinguishing of BCG type strains from M. bovis strains using a minute quantity of starting material. M. africanum identification indicated a tuberculosis microepidemic in a defined area when repeated isolation was observed . Sporadic isolation of M. africanum strains has been reported in Europe and the United States, including outbreaks of multidrug-resistant (MDR) strains ,. In recent studies, variations in the reported prevalence of M. africanum among various African countries may also reflect difficulties in accurate identification of this species (Figure 2). M. microti, M. pinnipedii, M. caprae and M. canettii remain difficult to identify because of the extremely slow growth of these organisms, the difficulties with their identification under traditional bacteriological methods  and the fact that these recently described species have not been incorporated into current molecular identification schemes.
ETR-D spacer sequencing offers a new tool for the rapid and accurate identification of all MTC species in a single sequencing reaction without the need for expensive, time-consuming and potentially harmful polyphasic approaches. Its use could assist public health interventions and aid in the control of zoonotic transmission in African countries. Accurate identification of MTC isolates from Africa and tropical Asia would be of particular interest from the perspective of the current emergence of multidrug resistant and extended resistance isolates in these countries .
The authors acknowledge the technical expertise of Christian Fontaine and Bernard Campagna. In addition, the authors recognize Dr. Christophe Guilhot, Dr. Sylvain Godreuil, and Dr. Cristina Gutierrez for providing the M. canettii clinical isolates.
Conceived and designed the experiments: Z. Djelouadji M. Drancourt. Performed the experiments: Z. Djelouadji. Analyzed the data: Z. Djelouadji D. Raoult M. Daffé M. Drancourt. Contributed reagents/materials/analysis tools: D. Raoult. Wrote the paper: Z. Djelouadji D. Raoult M. Drancourt.
- 1. Shah NS, Wright A, Bai GH, Barrera L, Boulahbal F, et al. (2007) Worldwide emergence of extensively drug-resistant tuberculosis. Emerg Infect Dis 13: 380–387.
- 2. Michel AL (2002) Implications of tuberculosis in African wildlife and livestock. Ann N Y Acad Sci 969: 251–255.
- 3. Cosivi O, Grange JM, Daborn CJ, Raviglione MC, Fujikura T, et al. (1998) Zoonotic tuberculosis due to Mycobacterium bovis in developing countries. Emerg Infect Dis 4: 59–70.
- 4. Frothingham R, Strickland PL, Bretzel G, Ramaswamy S, Musser JM, et al. (1999) Phenotypic and genotypic characterization of Mycobacterium africanum isolates from West Africa. J Clin Microbiol 37: 1921–1926.
- 5. Godreuil S, Torrea G, Terru D, Chevenet F, Diagbouga S, et al. (2007) First molecular epidemiology study of Mycobacterium tuberculosis in Burkina Faso. J Clin Microbiol 45: 921–927.
- 6. Niobe-Eyangoh SN, Kuaban C, Sorlin P, Cunin P, Thonnon J, et al. (2003) Genetic biodiversity of Mycobacterium tuberculosis complex strains from patients with pulmonary tuberculosis in Cameroon. J Clin Microbiol 41: 2547–2553.
- 7. van Soolingen D, Hoogenboezem T, de Haas PE, Hermans PW, Koedam MA, et al. (1997) A novel pathogenic taxon of the Mycobacterium tuberculosis complex, Canetti: characterization of an exceptional isolate from Africa. Int J Syst Bacteriol 47: 1236–1245.
- 8. WELLS AQ (1953) Mycobacterium tuberculosis var. muris. J Gen Microbiol 9: 149.
- 9. Mostowy S, Cousins D, Behr MA (2004) Genomic interrogation of the dassie bacillus reveals it as a unique RD1 mutant within the Mycobacterium tuberculosis complex. J Bacteriol 186: 104–109.
- 10. Cousins DV, Peet RL, Gaynor WT, Williams SN, Gow BL (1994) Tuberculosis in imported hyrax (Procavia capensis) caused by an unusual variant belonging to the Mycobacterium tuberculosis complex. Vet Microbiol 42: 135–145.
- 11. van Soolingen D, van der Zanden AG, de Haas PE, Noordhoek GT, Kiers A, et al. (1998) Diagnosis of Mycobacterium microti infections among humans by using novel genetic markers. J Clin Microbiol 36: 1840–1845.
- 12. Aranaz A, Liebana E, Gomez-Mampaso E, Galan JC, Cousins D, et al. (1999) Mycobacterium tuberculosis subsp. caprae subsp. nov.: a taxonomic study of a new member of the Mycobacterium tuberculosis complex isolated from goats in Spain. Int J Syst Bacteriol 49 Pt 3: 1263–1273.
- 13. Aranaz A, Cousins D, Mateos A, Dominguez L (2003) Elevation of Mycobacterium tuberculosis subsp. caprae Aranaz et al. 1999 to species rank as Mycobacterium caprae comb. nov., sp. nov. Int J Syst Evol Microbiol 53: 1785–1789.
- 14. Blaas SH, Bohm S, Martin G, Erler W, Gluck T, et al. (2003) Pericarditis as primary manifestation of Mycobacterium bovis SSP. caprae infection. Diagn Microbiol Infect Dis 47: 431–433.
- 15. Cousins DV, Bastida R, Cataldi A, Quse V, Redrobe S, et al. (2003) Tuberculosis in seals caused by a novel member of the Mycobacterium tuberculosis complex: Mycobacterium pinnipedii sp. nov. Int J Syst Evol Microbiol 53: 1305–1314.
- 16. van Soolingen D, de Haas PE, Haagsma J, Eger T, Hermans PW, et al. (1994) Use of various genetic markers in differentiation of Mycobacterium bovis strains from animals and humans and for studying epidemiology of bovine tuberculosis. J Clin Microbiol 32: 2425–2433.
- 17. Evans JT, Smith EG, Banerjee A, Smith RM, Dale J, et al. (2007) Cluster of human tuberculosis caused by Mycobacterium bovis: evidence for person-to-person transmission in the UK. Lancet 369: 1270–1276.
- 18. Witebsky FG, Kruczak-Filipov P (1996) Identification of mycobacteria by conventional methods. Clin Lab Med 16: 569–601.
- 19. Yates MD, Grange JM, Collins CH (1984) A study of the relationship between the resistance of Mycobacterium tuberculosis to isonicotinic acid hydrazide (isoniazid) and to thiophen-2-carboxylic acid hydrazide. Tubercle 65: 295–299.
- 20. Godreuil S, Torrea G, Terru D, Chevenet F, Diagbouga S, et al. (2007) First molecular epidemiology study of Mycobacterium tuberculosis in Burkina Faso. J Clin Microbiol 45: 921–927.
- 21. Cadmus S, Palmer S, Okker M, Dale J, Gover K, et al. (2006) Molecular analysis of human and bovine tubercle bacilli from a local setting in Nigeria. J Clin Microbiol 44: 29–34.
- 22. Kallenius G, Koivula T, Ghebremichael S, Hoffner SE, Norberg R, et al. (1999) Evolution and clonal traits of Mycobacterium tuberculosis complex in Guinea-Bissau. J Clin Microbiol 37: 3872–3878.
- 23. Kirschner P, Springer B, Vogel U, Meier A, Wrede A, et al. (1993) Genotypic identification of mycobacteria by nucleic acid sequence determination: report of a 2-year experience in a clinical laboratory. J Clin Microbiol 31: 2882–2889.
- 24. Frothingham R, Hills HG, Wilson KH (1994) Extensive DNA sequence conservation throughout the Mycobacterium tuberculosis complex. J Clin Microbiol 32: 1639–1643.
- 25. Woods GL (2001) Molecular techniques in mycobacterial detection. Arch Pathol Lab Med 125: 122–126.
- 26. Richter E, Weizenegger M, Fahr AM, Rusch-Gerdes S (2004) Usefulness of the GenoType MTBC assay for differentiating species of the Mycobacterium tuberculosis complex in cultures obtained from clinical specimens. J Clin Microbiol 42: 4303–4306.
- 27. Parsons LM, Brosch R, Cole ST, Somoskovi A, Loder A, et al. (2002) Rapid and simple approach for identification of Mycobacterium tuberculosis complex isolates by PCR-based genomic deletion analysis. J Clin Microbiol 40: 2339–2345.
- 28. Scorpio A, Zhang Y (1996) Mutations in pncA, a gene encoding pyrazinamidase/nicotinamidase, cause resistance to the antituberculous drug pyrazinamide in tubercle bacillus. Nat Med 2: 662–667.
- 29. Sreevatsan S, Escalante P, Pan X, Gillies DA, Siddiqui S, et al. (1996) Identification of a polymorphic nucleotide in oxyR specific for Mycobacterium bovis. J Clin Microbiol 34: 2007–2010.
- 30. Liebana E, Aranaz A, Francis B, Cousins D (1996) Assessment of genetic markers for species differentiation within the Mycobacterium tuberculosis complex. J Clin Microbiol 34: 933–938.
- 31. Prabhakar S, Mishra A, Singhal A, Katoch VM, Thakral SS, et al. (2004) Use of the hupB gene encoding a histone-like protein of Mycobacterium tuberculosis as a target for detection and differentiation of M. tuberculosis and M. bovis. J Clin Microbiol 42: 2724–2732.
- 32. Frothingham R (1995) Differentiation of strains in Mycobacterium tuberculosis complex by DNA sequence polymorphisms, including rapid identification of M. bovis BCG. J Clin Microbiol 33: 840–844.
- 33. Niemann S, Harmsen D, Rusch-Gerdes S, Richter E (2000) Differentiation of clinical Mycobacterium tuberculosis complex isolates by gyrB DNA sequence polymorphism analysis. J Clin Microbiol 38: 3231–3234.
- 34. Chimara E, Ferrazoli L, Leao SC (2004) Mycobacterium tuberculosis complex differentiation using gyrB-restriction fragment length polymorphism analysis. Mem Inst Oswaldo Cruz 99: 745–748.
- 35. Arnold C, Westland L, Mowat G, Underwood A, Magee J, et al. (2005) Single-nucleotide polymorphism-based differentiation and drug resistance detection in Mycobacterium tuberculosis from isolates or directly from sputum. Clin Microbiol Infect 11: 122–130.
- 36. Richter E, Weizenegger M, Rusch-Gerdes S, Niemann S (2003) Evaluation of genotype MTBC assay for differentiation of clinical Mycobacterium tuberculosis complex isolates. J Clin Microbiol 41: 2672–2675.
- 37. Viana-Niero C, Gutierrez C, Sola C, Filliol I, Boulahbal F, et al. (2001) Genetic diversity of Mycobacterium africanum clinical isolates based on IS6110-restriction fragment length polymorphism analysis, spoligotyping, and variable number of tandem DNA repeats. J Clin Microbiol 39: 57–65.
- 38. Ayele WY, Neill SD, Zinsstag J, Weiss MG, Pavlik I (2004) Bovine tuberculosis: an old disease but a new threat to Africa. Int J Tuberc Lung Dis 8: 924–937.
- 39. Frothingham R, Meeker-O'Connell WA (1998) Genetic diversity in the Mycobacterium tuberculosis complex based on variable numbers of tandem DNA repeats. Microbiology 144 ( Pt 5): 1189–1196.
- 40. Supply P, Magdalena J, Himpens S, Locht C (1997) Identification of novel intergenic repetitive units in a mycobacterial two-component system operon. Mol Microbiol 26: 991–1003.
- 41. Magdalena J, Supply P, Locht C (1998) Specific differentiation between Mycobacterium bovis BCG and virulent strains of the Mycobacterium tuberculosis complex. J Clin Microbiol 36: 2471–2476.
- 42. Adekambi T, Colson P, Drancourt M (2003) rpoB-based identification of nonpigmented and late-pigmenting rapidly growing mycobacteria. J Clin Microbiol 41: 5699–5708.
- 43. Niemann S, Richter E, Rusch-Gerdes S (2000) Differentiation among members of the Mycobacterium tuberculosis complex by molecular and biochemical features: evidence for two pyrazinamide-susceptible subtypes of M. bovis. J Clin Microbiol 38: 152–157.
- 44. Djelouagji Z, Drancourt M (2006) Inactivation of cultured Mycobacterium tuberculosis organisms prior to DNA extraction. J Clin Microbiol 44: 1594–1595.
- 45. Goh KS, Legrand E, Sola C, Rastogi N (2001) Rapid differentiation of “Mycobacterium canettii” from other Mycobacterium tuberculosis complex organisms by PCR-restriction analysis of the hsp65 gene. J Clin Microbiol 39: 3705–3708.
- 46. Jiang X, Zhang Y, Zhang W, Gao F, Jin J, et al. (2007) Identification of unique genetic markers in Rv0927c among Mycobacterium tuberculosis W-Beijing strains. Microbes Infect 9: 241–246.
- 47. Somoskovi A, Dormandy J, Parsons LM, Kaswa M, Goh KS, et al. (2007) Sequencing of the pncA gene in members of the Mycobacterium tuberculosis complex has important diagnostic applications: Identification of a species-specific pncA mutation in “Mycobacterium canettii” and the reliable and rapid predictor of pyrazinamide resistance. J Clin Microbiol 45: 595–599.
- 48. Huard RC, de Oliveira Lazzarini LC, Butler WR, van Soolingen D, Ho JL (2003) PCR-based method to differentiate the subspecies of the Mycobacterium tuberculosis complex on the basis of genomic deletions. J Clin Microbiol 41: 1637–1650.
- 49. Magdalena J, Vachee A, Supply P, Locht C (1998) Identification of a new DNA region specific for members of Mycobacterium tuberculosis complex. J Clin Microbiol 36: 937–943.
- 50. Sola C, Rastogi N, Gutierrez MC, Vincent V, Brosch R, et al. (2003) Is Mycobacterium africanum subtype II (Uganda I and Uganda II) a genetically well-defined subspecies of the Mycobacterium tuberculosis complex? J Clin Microbiol 41: 1345–1346.
- 51. Niemann S, Rusch-Gerdes S, Joloba ML, Whalen CC, Guwatudde D, et al. (2002) Mycobacterium africanum subtype II is associated with two distinct genotypes and is a major cause of human tuberculosis in Kampala, Uganda. J Clin Microbiol 40: 3398–3405.
- 52. Brudey K, Gutierrez MC, Vincent V, Parsons LM, Salfinger M, et al. C (2004) Mycobacterium africanum genotyping using novel spacer oligonucleotides in the direct repeat locus. J Clin Microbiol 42: 5053–5057.
- 53. Gutacker MM, Mathema B, Soini H, Shashkina E, Kreiswirth BN, et al. (2006) Single-nucleotide polymorphism-based population genetic analysis of Mycobacterium tuberculosis strains from 4 geographic sites. J Infect Dis 193: 121–128.
- 54. Daborn CJ, Grange JM, Kazwala RR (1996) The bovine tuberculosis cycle–an African perspective. Soc Appl Bacteriol Symp Ser 25: 27S–32S.
- 55. Gibson AL, Hewinson G, Goodchild T, Watt B, Story A, et al. (2004) Molecular epidemiology of disease due to Mycobacterium bovis in humans in the United Kingdom. J Clin Microbiol 42: 431–434.
- 56. Palenque E, Villena V, Rebollo MJ, Jimenez MS, Samper S (1998) Transmission of multidrug-resistant Mycobacterium bovis to an immunocompetent patient. Clin Infect Dis 26: 995–996.
- 57. Ahmed N, Alam M, Rao KR, Kauser F, Kumar NA, et al. (2004) Molecular genotyping of a large, multicentric collection of tubercle bacilli indicates geographical partitioning of strain variation and has implications for global epidemiology of Mycobacterium tuberculosis. J Clin Microbiol 42: 3240–3247.
- 58. Behr MA, Wilson MA, Gill WP, Salamon H, Schoolnik GK, et al. (1999) Comparative genomics of BCG vaccines by whole-genome DNA microarray. Science 284: 1520–1523.
- 59. Centers for disease control and prevention (CDC) (1997) Multiple misdiagnoses of tuberculosis resulting from laboratory error Wisconsin, 1996. MMWR Morb Mortal Wkly Rep 46: 797–801.
- 60. Desmond E, Ahmed AT, Probert WS, Ely J, Jang Y, et al. (2004) Mycobacterium africanum cases, California. Emerg Infect Dis 10: 921–923.
- 61. Kremer K, van Soolingen D, Frothingham R, Haas WH, Hermans PW, et al. (1999) Comparison of methods based on different molecular epidemiological markers for typing of Mycobacterium tuberculosis complex strains: interlaboratory study of discriminatory power and reproducibility. J Clin Microbiol 37: 2607–2618.
- 62. Cousins DV, Williams SN, Reuter R, Forshaw D, Chadwick B, et al. (1993) Tuberculosis in wild seals and characterisation of the seal bacillus. Aust Vet J 70: 92–97.
- 63. Shah NS, Wright A, Bai GH, Barrera L, Boulahbal F, et al. (2007) Worldwide emergence of extensively drug-resistant tuberculosis. Emerg Infect Dis 13: 380–387.
- 64. Kazwala RR, Daborn CJ, Sharp JM, Kambarage DM, Jiwa SF, et al. (2001) Isolation of Mycobacterium bovis from human cases of cervical adenitis in Tanzania: a cause for concern ?. Int J Tuberc Lung Dis 5: 87–91.
- 65. Kidane D, Olobo JO, Habte A, Negesse Y, Aseffa A, et al. (2002) Identification of the causative organism of tuberculous lymphadenitis in ethiopia by PCR. J Clin Microbiol 40: 4230–4234.
- 66. Koeck JL, Bernatas JJ, Gerome P, Fabre M, Houmed A, et al. (2002) Epidemiology of resistance to antituberculosis drugs in Mycobacterium tuberculosis complex strains isolated from adenopathies in Djibouti. Prospective study carried out in 1999. Med Trop 62: 70–72.
- 67. Mawak J, Gomwalk N, Bello C, Kandakai-Olukemi Y (2006) Human pulmonary infections with bovine and environment mycobacteria (atypical) in jos, Nigeria. Ghana Med J 40: 132–136.
- 68. Niemann S, Rusch-Gerdes S, Joloba ML, Whalen CC, Guwatudde D, et al. (2002) Mycobacterium africanum subtype II is associated with two distinct genotypes and is a major cause of human tuberculosis in Kampala, Uganda. J Clin Microbiol 40: 3398–3405.
- 69. Niemann S, Kubica T, Bange FC, Adjei O, Browne EN, et al. (2004) The species Mycobacterium africanum in the light of new molecular markers. J Clin Microbiol 42: 3958–3962.
- 70. Niobe-Eyangoh SN, Kuaban C, Sorlin P, Cunin P, Thonnon J, et al. (2003) Genetic biodiversity of Mycobacterium tuberculosis complex strains from patients with pulmonary tuberculosis in Cameroon. J Clin Microbiol 41: 2547–2553.