The genome of Mycobacterium tuberculosis and resistance mutations

1.   Types of tuberculosis and its therapy

1.1 Types and global incidence

One-third of the world’s population is thought to be infected with the tuberculosis (TB) bacillus [1]. Despite the availability of highly efficacious treatment for decades, TB remains a major global health problem. In 1993, the World Health Organization (WHO) declared TB to be a global public health emergency, at a time when an estimated 7–8 million cases and 1.3–1.6 million deaths occurred each year. In 2010, there were approximately 8.5–9.2 million cases and 1.2–1.5 million deaths from TB worldwide [2]. TB is the second leading cause of death from an infectious disease (after HIV, which caused an estimated 1.8 million deaths in 2008).

Global prospects for TB control are challenged by the emergence of mono-resistant strains, multidrug resistant and extensively drug resistant strains.Mono-resistance has been encountered against all anti-TB drugs and it arises by chromosomal mutation in a small proportion of bacilli in any wild strain. These resistant strains are selected by mono-therapy in conditions where the bacterial population is sufficiently large, as in cavitatory pulmonary disease.

Multidrug resistant (MDR) tuberculosis is caused by strains which are generally considered to be resistant to at least two drugs, such as isoniazid and rifampin. The term for extensively drug resistant (XDR) tuberculosis appeared in the literature in 2006 to describe a severe form of disease, presently defined as MDR-TB with additional bacillary resistance to any fluoroquinolone and at least one of three second-line injectable drugs: capreomycin, kanamycin and amikacin. XDR-TB is not yet the final product in the treatment of TB; amplification of drug resistance is still likely to occur if we do not manage XDR-TB properly [3].

There is also important to differentiate between primary and acquired resistance which is usually more severe. Primary resistance occurs in persons who have not received any anti-TB therapy and they are initially infected with drug resistant strains. Acquired resistance occurs in patients who have previously received anti-TB therapy and resistance develops as a result of inadequate regimen. Also, drug resistance among previously treated cases may not be truly acquired resistance, but contains a combination of several types of resistance: patients who have acquired resistance during TB treatment, patients who have been primarily infected with a resistant strain and subsequently failed therapy and patients who have been reinfected with a resistant strain.

The most recent estimates on the prevalence of anti-TB drug resistance come from surveys conducted by the WHO and the International Union Against Tuberculosis and Lung Disease [4]. These organizations investigated both new and previously treated TB cases in 93 geographical settings between 2002–2006. In these surveys, the prevalence of MDR-TB ranged from 0% to 22% among newly diagnosed cases and from 0% to 60% among previously treated cases. Rates of MDR-TB are especially high in some countries, including the former Soviet Union [1] and the frequency of acquired resistance to multi drugs is more common than primary resistance. In addition, since 2002, 45 countries have reported cases of XDR-TB. Of the MDR isolates tested for second-line drugs, 0%–30 % were found to be XDR.

Findings from modeling exercise warn that if MDR-TB case detection and treatment rates increase to the WHO target of 70%, without simultaneously increasing MDR-TB cure rates, XDR-TB could increase exponentially [3]. Clearly, control of drug resistant TB relies on preventing the emergence and amplification of drug resistance as well as timely diagnosis and proper management of drug-resistant disease.

1.2 Tuberculosis treatment

The treatment of tuberculosis faces three problems: (1) interruption of further transmission, (2) curing of the acute disease, (3) preventing relapse.

Anti-TB drugs were first introduced for TB therapy in the 1940s. However, single-drug therapy resulted in the rapid emergence of drug-resistant strains. By combining the 2 anti-TB drugs available at that time – streptomycin (STM) and para-aminosalicylic acid – the emergence of resistance was reduced to approximately 10%. When isoniazid (INH) was introduced in the early 1950s, and combined with STM and para-aminosalicylic acid, this combination effectively prevented the emergence of resistance but required 18 months of treatment to ensure cure of the disease. Subsequently, pyrazinamide (PZA), rifampicin (RIF) and ethambutol (EMB) were introduced and included together with isoniazid (INH) into the list of first-line drugs.

Extensive studies were carried out by the British Medical Research Council to define the optimal drug combination and the minimal duration of therapy[5]. The outcome consisted of a therapeutic regimen comprised of an initial 2-month treatment with INH, RIF and PZA, followed by a 4-month treatment with INH and RIF. Combination therapy is necessary for the treatment of the disease and to avoid the emergence of resistance, while a minimum 6-month treatment is required to prevent relapse. This protocol, further developed and now termed standard therapy short course, is still in use today and is recommended with slight modifications (including the addition of EMB or STM) by the International Union against Tuberculosis and Lung Disease, the World Health Organization and the American Thoracic Society [6].

To cure MDR-TB, healthcare providers must turn to a combination of second-line drugs, several of which are listed in Table 1. Second-line drugs may have more side effects, require up to 2 years of multidrug treatment, and the cost may be up to 100 times more than first-line therapy [7].

Table 1. Overview of anti-TB drugs: mechanisms of action and drug targets.

Drug

Cellular function inhibited

Target

First-line drugs
Isoniazid (INH) Mycolic acid synthesis Enoyl reductase
Rifampicin (RIF) RNA synthesis RNA polymerase
Ethambutol (EMB) Arabinogalactan synthesis Arabinosyl transferase
Pyrazinamide (PZA) Unclear Unclear
Second-line drugs
Fluoroquinolone (FLQ) DNA supercoiling DNA gyrase
Ethionamide (ETH) Mycolic acid synthesis Enoyl reductase
Streptomycin  (STM) Protein synthesis 30S ribosomal subunit
Kanamycin (KAN), Amikacin (AMK) Protein synthesis 30S ribosomal subunit
Capreomycin (CAP) Protein synthesis 30S/50S ribosomal subunit

Data from a large retrospective cohort (5550 patients) have provided evidence that standard shortcourse chemotherapy, based on first-line drugs, was inadequate to treat patients with MDR-TB [8]. A study from Russia demonstrated a high relapse rate (27.8%) of MDR-TB cases declared ‘successfully’ cured with standard short-course chemotherapy regimens (WHO category 1, e.g. 2 months with four drugs, INH-RMPEMB-PZA followed by 4 months with two drugs, INHRMP; and WHO category 2, e.g. 2 months with five drugs, INH-RMP-EMB-PZA-SM plus 1 months with four drugs INH-RMP-EMB-PZA followed by continuation phase of 5 months with three drugs, INH-RMPEMB) within a median time of 8 months (2.46 recurrences were observed in 100 person-months)[9]. Clearly, the category 2 retreatment regimen 2(INH-EMB-RMP-PZA-SM)/1(INH-EMB-RMP-PZA)/5(INH-EMB-RMP) recommended by WHO is not adequate for settings with a high proportion of MDR-TB patients among failure of category 1 regimen [10].

Treatment of XDR-TB is long and most complicated because these strains are resistant to the two best antibiotics, INH and RIF, as well as most of the alternative drugs used against MDR. It relies on drugs that are less potent, need to be administered for a much longer time and are substantially more toxic than those used to treat TB caused by drug-susceptible strains. Moreover, the cost of a second-line drug regimen is rather high: up to thousands of dollars compared with the cost of about $US 20 per patient for the standard 6-month short-course, first-line chemotherapy regimen (WHO category 1) [3].

 2.   Genome structure of M. tuberculosis  

2.1 The Mycobacterium tuberculosis complex

In 1882, in a remarkable feat of microbiology, Robert Koch isolated M. tuberculosis for the first time [11], and conclusively demonstrated in the guinea pig that this slow-growing mycobacterium was the agent of a human disease. Together with other highly related bacteria, M. tuberculosis forms a tightly knit complex, a single species as defined by DNA/DNA hybridization studies, which is characterized by a singular lack of diversity in the bulk of its genes. The M. tuberculosis complex (MTC) comprises six members (see Table 2): M. tuberculosis, the causative agent in the vast majority of human tuberculosis cases; Mycobacterium africanum, an agent of human tuberculosis in sub-Saharan Africa; Mycobacterium microti, the agent of tuberculosis in voles; Mycobacterium bovis, which infects a very wide variety of mammalian species including humans, and BCG (bacille Calmette–Guérin), an attenuated variant of M. bovis; and Mycobacterium canettii, a smooth variant that is very rarely encountered but causes human disease. Prior to the introduction of pasteurization of milk, M. bovis was responsible for ∼6% of total tuberculosis deaths in humans in Europe.

Table 2. Some properties of tubercle bacilli [11].

Species Host range Morphotype
Mycobacterium africanum Humans Rugose, dysgenic
M. bovis Badgers, cattle, deer, elephants, goats, humans, lions, sels, etc. Smooth, dysgenic
M. bovis BCG Rugose, cugonic
Mycobacterium canettii Humans Very smooth
Mycobacterium microti Voles Tiny
M. tuberculosis Humans Rugose, cugonic

BCG was derived by Calmette and Guérin from a virulent M. bovis isolate by 230 serial passages in a broth containing glycerol, potato-extract and bile salts. During the course of these passages the M. bovis strain progressively lost its virulence for animals and was first shown to be harmless and protective in a child in 1921. Since that time BCG has been used extensively as a live vaccine against tuberculosis and also protects humans against leprosy. Three billion doses have now been administered with negligible side effects and this is strong testimony to the safety of the vaccine. The attenuation process undergone by BCG probably involved the serial loss of genetic material, rendering reversion to virulence impossible. M. microti, the vole bacillus, is naturally attenuated for humans and has also been used successfully to protect against.

2.2 Genome structure

Genome of Mycobacterium tuberculosis has been sequenced with hopes of gaining further understanding of how to defeat the infamously successful pathogens.

The widely used reference strain M. tuberculosis H37Rv was firstly sequenced in 1998 [12]. Unlike some clinical isolates that often lose virulence after laboratory passaging this strain has retained full virulence in animals since its isolation in 1905.

Genome size of M. tuberculosis  is 4,411,522 base pairs long with  ∼4000 genes distributed fairly evenly between the two strands and accounting for >91% of the potential coding capacity, 6 pseudogenes, and a relatively high G+C content of 65.6%. At 4.4 Mbp, M. tuberculosis is one of the largest known bacterial genomes, coming in just short of E. coli, and a distant third to Streptomyces coelicolor [13].

Genes were classified into 11 broad functional groups and, today, precise or putative functions can be attributed to 52%, with the remaining 48% being conserved hypotheticals or unknown [14]. Over 51% of the genes have arisen as a result of gene duplication or domain shuffling events, and 3-4% of the genome is composed of insertion sequences (IS) and prophages (phiRv1, phiRv2). There are 56 copies of IS elements belonging to the well-known IS3, IS5, IS21, IS30, IS110, IS256 and ISL3 families, as well as a new IS family, IS1535, that appears to employ a frameshifting mechanism to produce its transposase. IS6110, a member of the IS3 family, is the most abundant element and has played an important role in genome plasticity [11].

About 250 genes are involved in fatty acid metabolism, with 39 of these involved in the polyketide metabolism generating the waxy coat. Such large numbers of conserved genes show the evolutionary importance of the waxy coat to pathogen survival. Approximately 10% of the coding capacity is taken up by two clustered gene families that encode acidic, glycine-rich proteins. These proteins have a conserved N-terminal motif, deletion of which impairs growth in macrophages and granulomas [15].

Unlike most bacteria which have multiple copies of the rRNA genes, M. tuberculosis contains a single rrn operon. This explains why single mutations in the 16S and 23S ribosomal RNA genes result in resistance to protein synthesis inhibitors and are very difficult to isolate [16].

3. Genetic diversity and geographical distribution

3.1 Mycobacterium strains genotyping methods

Historically, the microbiological tools used to differentiate or subspeciate clinical isolates of Mycobacterium tuberculosis were based on phage and/or drug susceptibility patterns. The comparison of strains has evolved from analysis of protein products (phenotyping) to the analysis of genetic content (genotyping).

TB genotyping is a laboratory-based approach used to analyze DNA of Mycobacterium tuberculosis due to specific sections of the M. tuberculosis genome that form distinct genetic patterns which help to distinguish different strains of M. tuberculosis [17]. Genotyping is important in research of genes and gene variants associated with disease. Genotyping results combined with epidemiologic data are used in a wide range of epidemiological, clinical and basic studies: to demonstrate foci of transmission in cites, nosocomial outbreaks, and congregate settings; to study transmission among high-risk populations such as HIV-positive and homeless persons; to evaluate cross-contamination in the clinical laboratory; and to analyze sequence changes as they reflect rates of molecular evolution.

The first genotyping methods of M. tuberculosis were introduced in 1990. In the ensuing years, more than 30,000 strains have been analyzed and cataloged, and thousands of unique patterns have been identified. Current methods of genotyping include restriction fragment length polymorphism identification (RFLP) of genomic DNA, random amplified polymorphic detection (RAPD) of genomic DNA, amplified fragment length polymorphism detection (AFLPD), polymerase chain reaction (PCR), DNA sequencing, allele specific oligonucleotide (ASO) probes and DNA microarrays  hybridization. Some of the techniques used nowadays are characterized below [18].

Restriction fragment length polymorphism analysis

This method is based on analyzing the distribution (both in number and location) of the target sequence in the chromosome of M. tuberculosis complex strains. Total DNA from the sample strain is digested with a restriction enzyme that cuts outside of the element, resulting in a collection of DNA fragments. Among this collection are a finite number of fragments of various lengths containing the target sequence RFLP. The fragments are separated by size, and those fragments bearing the target sequence are identified by Southern blot analysis. Each strain contains a distinct pattern of different-sized bands marked by the target sequence, although some caveats must be invoked.

The most commonly used target sequence is the bacterial insertion element IS6110. This mobile genetic element is 1355 bp in size, has an imperfect 28 bp inverted repeat, and generates a 3- to 4-bp target duplication upon insertion. It is present in 0–30 copies in the M. tuberculosis complex genome. The laboratory strain H37Rv, whose genome has been sequenced, contains 17 copies of IS6110. Pulsed field gel electrophoretic analysis and physical mapping of IS6110 copies in the chromosome of strain H37Rv have provided evidence that these insertions are randomly dispersed around the chromosome. The genomic context of these elements is the subject of several recent studies and will be discussed below.

Figure 1.  Example of IS6110-based RFLP image. Isolates represented by lanes 3, 5, 6, 9, and 10 have the same pattern and were epidemiologically linked. Lane S shows the CDC molecular weight standard [17].

Other genetic markers used for RFLP analysis include a second insertion element, IS1081, first identified in M. bovis and shown to be present in members of the M. tuberculosis complex strains. Also useful are the repeated elements such as the DR (direct repeat) locus, MPTR (major polymorphic tandem repeats), and PGRS (polymorphic GC-rich repetitive sequence).

Polymerase Chain Reaction

There are a number of fingerprinting techniques that exploit polymerase chain reaction (PCR) amplification. Using nucleotide primers of sequence derived from IS6110, for example, yields products using template concentrations as low as 10 pg. The DNA sequence of these products reveals priming from one of the IS6110 elements at one end and nonspecific sites in the chromosome at the other. PCR with random primers has also been used to analyze clustered cases.

Single-Strand Conformational Polymorphism

The analysis of drug-resistant isolates of M. tuberculosis on the basis of comparing the sequence of resistance genes has proven to be a very precise genotyping tool. Single-strand conformational polymorphism (SSCP) has distinguished strains on the basis of differences in the rpoB region among rifampin-resistant strains and gyrA among quinolone-resistant strains. A more direct method uses PCR amplicons derived from drug-resistant isolates followed by automated sequencing. The finding that 21 rpoB genes sequenced from wild-type strains have identical base pair changes supports the notion that these strains are clonal and are derived from a common ancestor.

Spoligotyping (Spacer Oligotyping)

This technique is based on sequence variation within a specific region of the M. tuberculosis chromosome, the highly polymorphic DR (direct repeat) locus. Its structure is a series of 36-bp imperfectly matched DR elements interrupted by spacer sequences. The DR elements vary in size from 35 to 41 bp and the spacers are 5–7 bp. In addition, the number of spacers is variable: BCG has 49 DR spacers and M. tuberculosis (H37Rv) has 39. PCR is used to amplify the DR region from each strain, and the PCR products are hybridized to membranes containing hundreds of immobilized random oligonucleotides. Each strain has a characteristic series of spacer regions in its DR locus, which is then reflected in the patterns detected by spoligotyping. Although the sequences vary, strains can be grouped on the basis of their spoligotype pattern.

An advantage of this technique is that the first step is PCR amplification and can be applied directly to clinical samples without waiting for expansion of the culture. IS6110 fingerprinting becomes less differentiating as a strain’s copy number decreases. Clinical strains with fewer than five copies of IS6100 are generally analyzed by alternate means. Several direct comparisons of spoligotyping and IS6110-based RFLP indicate that the DR locus can discriminate among strains containing as few as one copy of IS6110. In addition, there was an occasional splitting of a group of high-copy-number strains by spoligotyping.

Figure 2. Example of spoligotype patterns of certain strains [17].

Variable Number Tandem Repeat

A recent technique analyzed genetic loci containing tandem repeat sequences. Eleven variable number tandem repeat (VNTR) loci in the M. tuberculosisgenome were examined. Two types of repeat loci were identified. Five of the loci were major polymorphic tandem repeat (MPTR) loci and contained 15-base-pair variable repeats. The remaining six loci were exact tandem repeat (ETR) loci and contained identical sequences in large adjacent repeats. Spacer regions do not interrupt these tandem repeat loci, as in the DR loci discussed above. Primers were designed that recognize the termini of the repeat sequences, and the 11 loci were analyzed by PCR. The number of repeats present in each locus determined the length of the PCR product. These lengths were then compared among 48 strains. Seven of the 11 tandem repeat loci showed discriminating polymorphisms. For example, 22 of 25 M. tuberculosisstrains and 5 of 23 M. bovis BCG strains had distinct allele profiles. This technique should be useful for both strain differentiation and evolutionary analyses.

Direct DNA Sequencing

Finally, the application of automated sequencing techniques has permitted the analysis of several genes from large numbers of clinical strains. In addition, the complete H37Rv genome sequence has been annotated and published. The circular chromosome contains 4,411,529 base pairs with 3,918 protein coding regions. Most of these genes contain recognizable sequences, unlike gene sequences from other organisms, suggesting the function of their gene products. However, approximately 600 of the genes are entirely unknown. The availability of this information has significantly increased the pace of tuberculosis research. The applications of this approach are discussed below.

3.2 Phylogenetic analysis

Members of the MTC are considered genetically monomorphic with a high level of genomic sequence similarity (>99.95%), limited horizontal gene transfer, and a clonal population structure [19]. Single nucleotide polymorphisms (SNPs) do occur in the genomes of members of the M. tuberculosis complex but at a relatively low level for a bacterium of 1 in every 2000–4000 bp, depending on the species. Some SNPs, like the point mutation in the pncA gene responsible for pyrazinamide resistance, result in phenotypic change but the majority seems to be silent. Consequently, InDels appear to be the most common means of generating diversity where most of the insertions result from transposition events, generally involving IS6110, or more rarely from gene duplication [11].

This apparent homogeneity led to the assumption that genetic diversity among MTC strains would not be of clinical significance. However, recent data based on molecular genotyping methods (i.e. IS6110 RFLP, spoligotyping, MIRU-VNTR typing) revealed a highly diverse population structure with at least six major geographically-associated lineages that can be further subdivided into well-defined genotypes [19].

Phylogenetic analyses of the MTC have delineated multiple distinct lineages broadly divided into “modern” (clade 1) and “ancient” (clade 2) strains illustrated in Figure 3. Although molecular genotyping provides a robust framework for understanding evolution and epidemiology of MTC, the functional impact of genetic diversity remains poorly characterized.

Figure 3. Radial neighbor-joining tree showing genetic and transcriptome diversity of M. tuberculosis complex (MTC) clinical isolates [19]. Three strains each from the 5 distinct lineages pathogenic to humans plus two sequenced reference strains (H37Rv and CDC1551) were chosen to represent the global diversity of MTC.

 

Cluster analyses of the SNP markers from global collection of M. tuberculosis isolates identified six deeply branching, phylogenetically distinct SNP cluster groups (Figure 4). Analysis of these cluster groups strongly associated with the geographical origin of the M. tuberculosis samples and human hosts suggested that M. tuberculosis first arose in the Indian subcontinent and spread worldwide through East Asia [20].

Figure 4. Distribution of the spoligotype clades on the SNP-based phylogeny of a global collection of M. tuberculosis isolates [20]. Each isolate is indicated by a dot, which is color coded according to the spoligotype clade assignment: CAS – Central Asian clade; EAI – East African-Indian clade; H – Haarlem clade; LAM – Latin American and Mediterranean clade; PINI – Mycobacterium pinnipedii clade; S  – S clade; T – T clade; X – X clade.

A mixed representation (by absolute case number and by frequency) of the distribution of the most frequent genotype families of M. tuberculosis is illustrated in Figures 5 and 6. They are grouped by genetic lineage, for the following six MTC lineages: Beijing, M. bovis, Central-Asia, East-African-Indian, Haarlem and Latin-American-Mediterranean. These Figures provides the best display of the global phylogeographical structure of the MTC population [21].

Figure 5. Synthesizing World Maps showing absolute (diameter) and percentage (colour) numbers of 3 genotype families within each country [21]: Beijing; EAI (East-African Indian) CAS (Central Asia).

Figure 6. Synthesizing World Maps showing absolute (diameter) and percentage (colour) numbers of 3 genotype families within each country [21]M. bovis; Haarlem; Latin-American and Mediterranean (LAM).

A special place occupies genetic family Beijing, first identified in histological preparations of lung tissue in the years 1956-1990 from patients of Beijing suburbs. In this group, the frequency of multidrug resistance is significantly higher than among other families. In brief, Beijing and Beijing-like strains represent about 50% of the strains in Far East-Asia and 13% of isolates globally. The Beijing genotype which may have been endemic in China for a long time is emerging in some parts of the world, especially in countries of the former Soviet Union, and to a lesser extent in the Western world [21].

4.   Molecular basis and genetic aspects of drug resistance

 4.1 Drug resistance mutations Mycobacterium tuberculosis

Drug resistance in TB is believed to be mediated exclusively by chromosomal mutations, which affect either the drug target itself or bacterial enzymes that activate prodrugs. Since the early 1990s, numerous studies have described the genetic mechanisms of drug resistance in Mycobacterium tuberculosis, and a wealth of data has accumulated on the mutations found in isolates resistant to specific drugs [22].

Several factors influence the degree of success of treatment programs including duration and complexity of therapy, ease of healthcare access, treatment cost, patient adherence, and drug side effects. Drug resistance in M. tuberculosis occurs when resistant mutants that are present naturally in the mycobacterial population are selected out by inadequate or interrupted treatment. Mutants resistant to a single drug occur approximately in every 10– 6 to 10– 8 cells. In theory, the effective presence of a mutant which is resistant to 2 drugs would require a population of 1012 – 1016 mycobacterial cells [6]. This mathematical concept provides the basis for the successful use of combination drug therapy to prevent the emergence of resistance.

It has been observed that drug-resistant mutants were less virulent as compared with the susceptible original strain [23] and that acquisition of drug resistance in bacteria is often accompanied by a “cost” of reduced bacterial growth. However, there is also low-cost or no-cost mutation.  It has been reported that drug-resistant strains of M. tuberculosis exhibit a wide range of virulence [24]. Furthermore, resistance mutations that incur an initial fitness cost may be compensated by later mutations that restore an organism’s reproductive potential [3].

Many studies suggest that MDR-TB strains primarily arise as a consequence of sequential accumulation of mutations in individual drug target genes. The development of resistance in TB begins with monoresistance, and subsequent resistance to additional drugs may occur. The order of mutation could vary widely, and may rarely begin with fluoroquinolone resistance [3]. A well documented example of how multi drug resistant Mycobacterium tuberculosis strains arise has been provided by the analysis of the evolution of two closely related subclones in New York City designated as a strain W and W1. These two related organisms have caused greater than 300 cases of tuberculosis in New York City and elsewhere. Automated DNA sequencing of representative organisms defined the exact series of distinct mutations conferring resistance to Rifampin, INH, Streptomycin, Ethambutol, ETH, PZA, Kanamycin, and quinolones [25].

In addition, resistance to a single drug may involve multiple genetic alterations locating to different genes, as well as multiple genetic alterations within a single gene. Presumably, this accumulation of various resistance mutations, all associated with resistance to a single drug, will either affect (increase) phenotypic resistance or ameliorate the fitness cost associated with a defined resistance mutation [6]. On the contrary, there has been no single genetic mutation identified resulting in resistance to two or more anti-TB drugs [26].

The target regions of M. tuberculosis and the mutations conferring drug resistance in patients with Tuberculosis are summarized in Table 3. Resistance mechanisms for the mostly used anti-tuberculous agents are summarized below.

Table 3. Target regions of Mycobacterium tuberculosis and the mutations conferring drug resistance in patients with Tuberculosis [4,27].

Drug

Line of therapy

Gene

Codon mutation sites

Rifampin (RIF)

1st

rpoB

513, 526, 531, 533

Isoniazid (INH)

1st

mabA-inhA,

katG,

oxyR-ahpC

8, 15

279, 315

39, 46

Ethambutol (EMB)

1st

embB

306, 406, 497

Pyrazinamide (PZA)

1st

pncA

63, 138, 141, 162

Streptomycin (STM)

1st

rpsL

rrs

43, 88

491, 512, 513, 516, 906

Ethionamide (ETH)

2nd

mabA-inhA

94

Fluoroquinolones (FLQ)

2nd

GyrA

74, 90, 91, 94, 102

Rifampin (RIF) is not specific for mycobacteria but affects many bacteria by interacting with the RNA polymerase β-subunit and preventing transcription[6]. The highly effective bactericidal action of this drug against M. tuberculosis has made it a key component of the initial anti-tuberculous regimen. Analysis of Rifampin strains from global sources has found that 96% of Rifampin resistant clinical isolates of Mycobacterium tuberculosis have mutations in the 81-bp core region of rpoB gene, which encodes the β-subunit of RNA polymerase [28,29]. These mutations are absent in susceptible organisms. Although minor discrepancies have been reported, in general there have been a strong correlation of a specific amino acid substitutions and MIC. Missense mutations in codons 513, 526, or 531 result in high level Rifampin resistance, whereas amino acid changes at position 514 or 533 usually result in low levels of Rifampin resistance. Also some studies has shown that laboratory derived Rifampin resistant mutants had a statistically significant fitness cost but this cost was less in rpoB S531L mutants [3].

It is estimated that 90% of rifampin-resistant isolates in some areas are also resistant to isoniazid, making rifampin resistance a useful surrogate marker for multidrug resistance and indicating that second and third line drugs to which these isolates are susceptible are urgently required.

Isoniazid (INH) is a synthetic, bactericidal agent, used as a first-line TB drug. INH is a prodrug which needs to be converted into its active form by the bacterial enzyme KatG, a catalase- peroxidase. Biochemical and genetic studies suggest that activated INH targets InhA, an enoyl-acyl carrier protein reductase, which takes part in fatty acid biosynthesis [6]. Investigators on several continents have reported that many (50-60%) INH resistant patient isolates have mutations, small deletions or insertions that are not represented among INH sensitive control strains [29]. Mutations leading to INH resistance have been identified in different gene targets including KatG, inhA, ahpC and other genes. Some researchers analyzed Mycobacterium tuberculosis isolates by PCR and found that the mutation frequencies were as follows for INH resistant strains KatG (36.8%), inhA (31.6%), KatG-inhA (2.6%), ahpC (13.2%) and KatG-ahpC (2.6%) [30].

Majority of resistant organisms has amino acid changes in KatG or nucleotide substitutions upstream of inhA. Varying by geographic area, 50 to 100% of INH resistant strains has mutations in codon 315 of the katG gene or in the promoter region of the inhA gene [31]. While mutational alterations in inhAmainly affect the gene’s promoter resulting in InhA overexpression and are associated with low-level resistance to INH, mutations in katG confer moderate- to high-level drug resistance [6]. A region of katG is also frequently altered in resistant strains with residue Arg463 [32]. However, mutations in the KatG or inhA do not account for all INH resistant strains since 15-25% INH resistance clinical isolates have both wild-type KatG and inhA genes. The mechanism of INH resistance in some strains remains to be determined [27].

Streptomycin (STM) is another first-line TB drug. Mutations associated with streptomycin resistance in tuberculosis have been identified in the 16S rRNA gene (rrs) and rpsL gene. In contrast to other bacteria that have multiple copies of rRNA genes, mycobacterium tuberculosis complex members have only one copy. Therefore, single nucleoside changes can potentially produce antibiotic resistance. Mutations in the rrs are clustered in two regions around nucleotides 530 and 951. The 530 loop 16S rRNA is highly conserved and is located adjacent to the 915 region in secondary structure models. The majority of mutations producing streptomycin resistance occur in rpsL. The most common mutation is at the codon 43. Mutations also have occurred in codon 88. About 65-75% of streptomycin resistant isolates have resistance-associated changes in rpsL or rrs. Failure to identify resistance-associated variations in these genes in 25-35% of organisms indicates that other molecular mechanisms of streptomycin resistance exist [27].

Pyrazinamide (PZA) is a structural analogue of nicotinamide that is used as a first-line TB drug. PZA kills semi-dormant tubercle bacilli under acidic conditions. It is believed that in the acidic environment of phagolysosomes the tubercle bacilli produce pyrazinamidase, an enzyme that converts PZA to pyrazinoic acid, the active derivative of this compound. To define the molecular mechanism of PZA resistance the pncA gene encoding pyrazinamidase has been sequenced. The results have provided evidence that pncA mutations conferred PZA resistance. DNA sequencing of PZA resistance clinical isolates identified mutations at codons, 63, 138, 141, and 162. In contrast, susceptible organisms had wild type sequences. Lack of pncA mutations in 28% of PZA resistant isolates suggested the existence of at least one additional gene participating in resistance. A remarkably wide array of pncA mutations resulting in structural changes in the PncA has been identified in greater than 70% of drug resistant isolates. It is presumed that these structural changes detrimentally change enzyme function, thereby altering conversion of PZA to its bioactive form [27].

The testing of PZA is problematic. PZA is a unique and unconventional antibiotic that is not active in vitro under normal culture conditions. The quantitative analysis of resistance levels, for example, low-level versus high-level resistance, is not possible with the procedures currently in place. However, as PZA is ineffective against Mycobacterium bovis, which is naturally pncA-, it has to be assumed that pncA negativity confers high-level drug resistance – at least in clinical terms [6].

Ethambutol (EMB) is a bactericidal first-line TB drug. This agent is thought to act on the mycobacterial cell wall, with arabinan synthesis as the primary site of action. Drug susceptibility testing for EMB is particularly problematic [6]. To understand the mechanism of resistance of Ethambutol a two gene locus (embAB) that encodes arabinosyl transfer has been established. Automated sequencing of these regions in clinical isolates from diverse geographical areas, discovered that 69% of Ethambutol resistance isolates had an amino acid substitution in EmbB that was not found in Ethambutol susceptible strains. The great majority (98%) of strains had mutations in codon 306, however, mutations were also identified in 3 additional codon 285, 330, and 630. These mutations were also uniquely represented among Ethambutol resistance organisms. The data are consistent with the idea that specific amino acid substitutions in EmbB detrimentally affect the interaction between Ethambutol, a putative anabinose analogue and EmbB likely to be an arabinosyl transferase. EmbB mutations are associated with Ethambutol resistance in roughly 70% of Ethambutol isolates of Mycobacterium tuberculosis. The cause of Ethambutol resistance in many organisms lacking mutations in ERDR of EmbB is unknown [27].

Ethionamide (ETH) is a second-line TB drug that is thought to inhibit mycolic acid biosynthesis in Mycobacterium tuberculosis. Studies have shown that for certain strains, low level of INH resistance is correlated with co-acquisition of Ethionamide resistance, suggesting that INH and Ethionamide share a common molecular target and most likely the mab-inhA genes [27].

Fluoroquinolones (FLQ) are a family of synthetic broad-spectrum antibiotics, which eradicate bacteria by interfering with DNA replication. Drug resistance in TB rarely begins with fluoroquinolone resistance. The history of TB treatment has observed sequential development of resistance to key anti-TB drugs over the decades, from SM resistance, to INH resistance, RMP resistance, then fluoroquinolone resistance. Unfortunately, international recommendations and national policies on protecting fluoroquinolone are not yet in place [3].

In general, there is a clear correlation between the genetic mechanism and the resistance phenotype. Thus, mutations in rpsL (STM), rpoB (RIF) or 16S ribosomal RNA (KAN, AMK, 2- deoxystreptamine aminoglycosides) are associated with high-level drug resistance, and mutations in gldB  (STM), eis(kanamycin, KAN), and inhA (INH) confer a low-level resistance phenotype. Resistance-conferring chromosomal alterations in a drug target gene are highly restricted. Presumably this reflects the in vivo selection for resistance mutations which maintain gene function, readily explaining the predominance of certain resistance mutations. In contrast, resistance-conferring chromosomal alterations in genes involved in prodrug conversion, for example pncA andethA, often display a wide diversity, indicating that there is little functional constraint as a loss of gene function phenotype is apparently well tolerated [6].

4.2 Geographical distribution and surveillance of mutations

Differences in the type and frequency of mutations in drug-resistant isolates in diverse geographical regions can pose challenges for the development of a sequence-based diagnostic, especially if the test relies on detecting a limited number of mutations. Different drugs of the same drug class may also interact differently with the molecular drug targets and thus affect the mutations selected for. In addition, some specific mutations have been shown to lead to high-level resistance, while others confer low-level resistance to a specific drug. Others may lead to cross-resistance to drug analogs. Regional differences in drug analogs used have been shown to select for different drug resistance mutations; for example, mutations found in regions that use specific rifamycin analogs differ from those in which other analogs are used. The mutations that arise from these different drug analog exposures may help explain some of the reported geographical differences in drug efficacy. We have therefore included the geographic sites of resistance studies in our database. As more data on the frequency of mutations become available through large-scale sequencing projects, we will garner an even more comprehensive understanding of the global distribution of specific mutations and a better perception of the role of geographical differences [4].

5.   Identification of drug resistant tuberculosis

5.1 Drug-Susceptibility Testing

During the 1950s, the establishment of laboratory methods for M. tuberculosis drug susceptibility testing (DST) was a tremendous challenge. At that time, when diagnostic procedures for drug susceptibility testing of bacteria were largely unexplored, sensitivity and resistance in M. tuberculosis were defined as follows: “sensitive” strains are those that have never been exposed to anti-TB drugs (wild strains); “resistant” strains are those that differ from sensitive strains in their capacity to grow in the presence of higher concentrations of the drug [33].

Nowadays, reliable and valid DST is essential for optimal DR-TB treatment. A number of different techniques are available for this and can be performed as direct or indirect tests. In the direct test, the test is performed with a directly inoculated concentrated specimen. An indirect test involves inoculation with a pure culture grown from the specimen.

Phenotypic methods involve culturing of M. tuberculosis in the presence of anti-TB drugs to detect inhibition of growth. Phenotypic methods allow the detection of drug resistance regardless of mechanism or molecular basis. However, phenotypic DST is a timeconsuming process because it requires culturing, which may take up to two months or longer. Genotypic approaches detect the genetic determinants of resistance rather than the resistance phenotype. Genetic testing is often referred to as “rapid molecular testing” since results can be available with less than one day turnaround time.

Many regional laboratories are able to perform susceptibility testing for only four first-line drugs: isoniazid, rifampicin, ethambutol, and streptomycin. Pyrazinamide-susceptibility testing is especially challenging and cannot be done using conventional solid media [34].

Susceptibility testing for second-line antituberculous drugs is not as simple as testing for isoniazid and rifampicin, partly because critical drug concentrations defining resistance are much closer to the minimal inhibitory concentrations in the former than in the latter. It is true that proficiency metrics equivalent to those obtained for first-line drug testing are not available for second-line drug testing Nonetheless, reasonably consistent, clinically validated criteria for drug resistance testing were determined almost 50 years ago, for the proportion method on Lowenstein-Jensen medium for kanamycin, capreomycin, viomycin, d-cycloserine, ethionamide, and PAS. Guidance on susceptibility testing for second-line drugs was published in 2001 by WHO [35,36]. Since that time new testing methods have been developed and additional standardization of existing methodologies has been undertaken. However, a validity study of second-line DST is currently underway.

Despite there has been much debate on the reliability (the ability of the results to be consistently reproducible) of second-line DST, these do not negate the usefulness of in vitro susceptibility testing for antituberculous drugs; many treatment facilities and programs have obtained adequate outcome results with regimens based on first- and second-line DST [33]. Any DST results should be interpreted with care, taking into consideration local resistance prevalence, the patient’s drug history, and the quality of the testing laboratory.

5.2 Phenotypic testing in the laboratory

Minimum inhibitory concentration (MIC) is the lowest concentration of an antimicrobial that will inhibit the visible growth of a microorganism after overnight incubation. Visibility of growth starts at 5×105 cells/ml. It was found that drug-susceptible strains of M. tuberculosis that have not been exposed to anti-TB drugs (wild-type strains) do not exhibit much variation in MICs to those drugs.

The critical concentration is defined as drugconcentration that inhibits the growth of wild-type strains, without appreciably affecting the growth of strains with alterations in drug susceptibility (see Table 4). This categorizes a clinical M. tuberculosis isolate as either susceptible or resistant [6].

Table 4. Mycobacterial drug succeptibility testing: the critical concentration (mg/l).

Antimicrobial agent

MIC of susceptible M.tuberculosis

Concentration (mg/l) in serum

Concentration used for testing (mg/l)

BACTEC 12B low

BACTEC 12B high

INH 0,05-0,2 5-10 0,1 0,4
RIF 0,5 10 1
PZA 20 40-50 100
EMB 1-5 2-5 5 7,5
Ofloxacin 0,25-0,5 2-10 2
Ethionamide 0,5-2,5 2-20 2,5
STM 0,5-1,0 25-50 2 6
AMK 0,5-1,0 20-40 1
Capreomycin 2-5 10-30 5

Standardization of the critical concentration was not been without controversy. For example, the recommended concentrations for EMB underwent adjustments over time, not the least because of low interlaboratory reproducibility of EMB susceptibility testing. Most likely, this is due to the very small differences between the concentration used for in vitro drug susceptibility testing and the natural drug susceptibility of wild-type isolates of M. tuberculosis. Thus, minute changes in drug susceptibility will have a major impact on interpretation of in vitro test results, with only a narrow range between the MICs of susceptible isolates and resistant isolates.

The proportion methods are based on observation that all strains of tuberculosis contain some bacilli that are resistant to antibacillary drugs – in resistant strains, the proportion of such bacilli is considerably higher than in sensitive strains (see Table 5).

Table 5. Critical concentration and critical proportion.

Antimicrobial agent Concentration used for testing (mg/l) Critical proportion, %
INH 0,2 1
RIF 4 1
PZA 100 10
EMB 2 10
STM 4 10

Critical proportion for resistance of a bacterial wild-type population follows a Gaussian distribution (see Figure 7). Thus, depending on the drug concentration, a small fraction of the population will show phenotypic resistance. This observation forms the basis for combining proportion testing and critical concentration.

Figure 7. Distribution of critical proportion for resistance depending on drug concentration.

Based on these assumptions, laboratory assays for INH and RIF, for example, compare the growth of a 1:100 dilution of the M. tuberculosis isolate on media without drug with growth of the undiluted suspension on media containing each drug. If the undiluted suspension grows faster or more abundantly in the presence of the drug than does the 1:100 dilution in the absence of the drug, the isolate is considered to contain a resistant population greater than 1% and is reported as resistant.

Among mycobacteriologists a misperception of critical proportion and clinical resistance frequently prevails. While the critical proportion of cells (subpopulation) able to grow in the presence of the critical concentration is mostly defined as equal or greater than 1% of the population [6], the frequency of mutational resistance is much lower, approximately 0.00001% of the population. It is, however, the mutational resistance which is responsible for treatment failure and for the emergence of resistance following inappropriate drug regimens. The critical proportion of resistance is a technical term and should not be confused with mutational resistance. In combination with the critical concentration, the critical proportion is a laboratory term used in in-vitro drug susceptibility testing to define the epidemiological cut-off (see Figures 8-10).

Genes involved in resistance

Role in resistance

Phenotypic resistance

Frequency in clinical strains

katG

Prodrug conservation

Moderate- to high-level (>1 mg/l)

70-80%

inhA

Drug tagret

Low-level (<1 mg/l)

20-30%

Figure 8. Schematized changes in drug susceptibility – exemplary Gaussian distributions of a population’s drug susceptibility for INH [6].

Genes involved in resistance

Role in resistance

Phenotypic resistance

Frequency in clinical strains

rpoB

Drug target

mostly high-level (>20 mg/l)

>95%

Figure 9. Schematized changes in drug susceptibility – exemplary Gaussian distributions of a population’s drug susceptibility for RIF [6].

Genes involved in resistance

Role in resistance

Phenotypic resistance

Frequency in clinical strains

embB

Drug target

Low- to moderate-level (<25 mg/l)

>80%

Figure 10. Schematized changes in drug susceptibility – exemplary Gaussian distributions of a population’s drug susceptibility for EMB [6].

5.3 Molecular diagnostics tests

Conventional tests for laboratory confirmation of TB include acid-fast bacilli (AFB) smear microscopy, which can produce results in 24 hours, and culture, which requires 2-6 weeks to produce results. Although rapid and inexpensive, AFB smear microscopy is limited by its poor sensitivity (45%-80% with culture-confirmed pulmonary TB cases) and its poor positive predictive value (50%-80%) for TB in settings in which nontuberculous mycobacteria are commonly isolated. Better efforts to control TB require faster and more accurate diagnostic tests rapid molecular diagnostics tests, which can give results in 3–6 hours, have been developed to address these issues [37].

Nucleic Acid Amplification Tests

Nucleic-acid amplification tests, also known as NATs or NAATs, are used to identify small amounts of DNA in test samples. There are several different kinds of nucleic-acid amplification tests, but they are all based on the same principal. A nucleic-acid amplification test uses a series of repeated reactions to make numerous copies of the DNA that someone is trying to detect. This amplifies the signal of the nucleic acids in the test sample so that they are easier to identify.

Once the amount of DNA has been increased in the sample using PCR or LCR, more conventional tests are used to detect it. These tests usually involve some form of nucleic acid hybridization, where the sample is probed with an artificially produced complementary strand of DNA that has been labeled in some way that makes it easy to detect and analyze.

Tests include those that are “in-house”, when they are based on a protocol developed in a non-commercial laboratory (“home-brew”), or commercial kits. Several commercial NAATs exist, and each uses a different method to amplify specific nucleic-acid regions in the Mycobacterium tuberculosis complex. These kits include: the GenProbe Amplified M. tuberculosis Direct test (AMTD), the Roche Amplicor MTB test, the Cobas Amplicor test, the Abbott LCx test, and the BD-ProbeTec (SDA) test [38].

According to WHO recommendations NAATs should be performed on at least one respiratory specimen from each patient with signs and symptoms of pulmonary TB for whom a diagnosis of TB is being considered but has not yet been established, and for whom the test result would alter case management or TB control activities. The following testing and interpretation algorithm is proposed.

Loop-Mediated Isothermal Amplification Tests

Molecular methods, in particular NAATs, have demonstrated high specificity and sensitivity approaching that of culture for the diagnosis of pulmonary TB. However, the cost and moderate complexity of NAATs have limited their widespread use, even in industrialized countries. It may be possible to dramatically simplify NAAT technology by using isothermal amplification methods with visual readout [33] and by engineering the sputum processing steps to provide a relatively inexpensive, handheld device. FIND is partnering with Eiken Chemical Co., Ltd., on the development of such a system using its LAMP (Loop-Mediated Isothermal Amplification) technology. This technology uses isothermal amplification, which removes the need for a thermocycler, in a closed tube, which reduces the chance of false-positive results from workspace contamination with amplified DNA. It is a very rapid reaction, with results available in less than 45 minutes, and, critically, generates results (based on turbidity and fluorescence) that can be seen with the naked eye. To date work has advanced through several prototypes with the current test system having only three plastic devices and a single instrument, a heating block. The performance of TB-LAMP prototypes suggests that the technology may have the potential to revolutionize diagnostic care at microscopy center.

5.4 Rapid Detection of Drug-Resistant TB Strains

WHO guidelines recommend that patients with a positive rapid molecular test be started on a standardized regimen for MDR-TB [39]. At the same time, sputum can be sent for phenotypic culture-based DST to additional first- and second-line drugs. When these more complete results are available, the treatment regimen may be modified accordingly. The use of rapid molecular testing in this way significantly reduces the time to effective treatment for MDR-TB patients, and also decreases nosocomial transmission by quickly identifying highly infectious MDR-TB patients.

In general, several options of rapid testing of anti-TB drug resistance are available [3], including DNA sequencing [40], solid-phase hybridization techniques [41], microarrays [42], real-time PCR techniques [43], microscopic observation drug susceptibility assay [44], slide DST [45], phage-based assays[46], colorimetric methods (the MTT and resazurin as redox indicators) [47] and nitrate reductase assay [48].

Molecular Assays are potentially the most rapid and sensitive methods for the detection of drug resistance and are theoretically able to provide a same-day diagnosis from clinical samples. The utility of these assays is dependent on their ability to detect all common drug resistance mutations.

All designed methods to exploit the observation that specific polymorphisms found in resistant strains are absent in susceptible organisms. The fact that natural populations of drugs-susceptible Mycobacterium tuberculosis isolates recovered globally have remarkably few polymorphisms in structural genes greatly simplifies interpretation of these assays. In essence, the M. tuberculosis complex is an ideal situation for application of certain kinds of molecular diagnostic testing strategies [27]. Each molecular strategy has advantages and disadvantages.

5.5 The line-probe assay for diagnosis of first-line drug resistance

The line-probe assay strategy has the advantage of relatively reliable performance, and potential commercial availability [27]. This is ideally suited for genotypic assays, since it is able to detect amplicons or PCR products as they are synthesized during real-time PCR and to discriminate between DNA sequences that differ from one another by as little as a single nucleotide substitution [49].

All strategies suffer from the fact that for no antimycobacterial agents do we understand the molecular of resistance for 100% of organisms. Hence, identification of a resistance-associated mutation is clinically informative whereas lack of a mutation in the target sequence must be interpreted with considerable caution.

Line-probe assay technology involves the following steps [50]:

–          First, DNA is extracted from M. tuberculosis isolates or directly from clinical specimens.

–          Next, polymerase chain reaction (PCR) amplification of the resistance-determining region of the gene under question is performed using biotinylated primers.

–          Following amplification, labeled PCR products are hybridized with specific oligonucleotide probes immobilized on a strip.

–          Captured labeled hybrids are detected by colorimetric development, enabling detection of the presence of M. tuberculosis complex, as well as the presence of wild-type and mutation probes for resistance. If a mutation is present in one of the target regions, the amplicon will not hybridize with the relevant probe. Mutations are therefore detected by lack of binding to wild-type probes, as well as by binding to specific probes for the most commonly occurring mutations. The posthybridization reaction leads to the development of coloured bands on the strip at the site of probe binding and is observed by eye.

Currently the two main line-probe diagnostic tests available commercially [4] are the INNO-LiPA TB test (Innogenetics) and the GenoType MTBDRplus kit (Hain Lifescience). These assays have recently been approved by the World Health Organization as a tool for rapid MDR-TB diagnosis. These assays are based on the detection of a set number of mutations in a few genes associated with resistance against isoniazid and rifampicin. These assays may therefore be unable to detect all isoniazid and rifampicin resistance and do not detect resistance to other first-line or second-line drugs.

INNO-LIPA TB assay

The INNO-LIPA Rif. TB assay (Innogenetics), is an LPA that uses multiplex PCR amplification and reverse hybridization to identify M. tuberculosis complex and mutations to the rpoB gene in culture isolates and has been available for more than 10 years. The INNO-LiPA Rif.TB kit contains 10 oligonucleotide probes, ie. one specific for the M. tuberculosis complex, five overlapping wild-type (susceptible) probes, and four probes for detecting specific mutations (D516V, H526Y, H526D and S531L) associated with rifampicin resistance, immobilized on nitrocellulose paper strips [50]. The region from codon 509 to 534 is covered by the wild-type probes.

Figure 11. Position of the oligonucleotide probes on the INNO-LiPA Rif.TB strip [38].

The INNO-LiPA Rif.TB assay has been labeled for use on isolates of M. tuberculosis complex grown on solid media. Information on expanded certification for use of INNO-LiPA Rif.TB on clinical specimens is pending.

In a large study of rapid tests for MDR-TB in Peru supported by FIND and WHO/TDR, this test also had excellent performance when applied directly to AFB smear-positive sputum specimens.

GenoType MTBDRplus assay

The GenoType® MTBDRplus assay (Hain Lifescience, Nehren, Germany) is a commercially available assay that combines detection of MTB complex with prediction of resistance to RIF and INH [31]. Genotype MTBDRplus, the second-generation assay, also detects mutations in the inhA gene that confers resistance to low-levels of isoniazid [50].

In the assay a multiplex PCR is followed by hybridization of the obtained DNA amplicons to membrane-bound probes. The assay combines detection of MTB complex with detection of mutations in the 81-bp hotspot region of rpoB, at codon 315 of the katG gene and in the inhA promoter region. For rifampicin resistance, four specific rpoB mutations are detected, ie. D516V, H526Y, H526D, and S531L. Eight wild-type probes are also present which cover the region from codon 505 to 533. For isoniazid resistance, S315T1 and S315T2 katG mutation probes are detected, together with four inhA mutation probes, i.e. C15T, A16G, T8C and T8A.

The Genotype MTBDRplus assay has been validated for use directly on smear-positive pulmonary specimens, as well as on isolates of M. tuberculosisgrown on solid medium or in liquid medium, after specimen processing by NaOH-NALC. A recent meta-analysis pooled all these studies and calculated pooled sensitivity and specificity rates of 99% (95% confidence interval (CI), 96%-100%) and 99% (95% CI, 98%-100%) respectively for rifampicin resistance, and of 96% (93-98%) and 100% (99-100%), respectively for isoniazid resistance.

Figure 12. Example of GenoType MTBDRplus strips [50].

In June 2008, the World Health Organization (WHO) endorsed the use of molecular line-probe assays for MDR-TB screening [31], and the GenoType® MTBDRplus assay has since been introduced for routine practice in various countries. The WHO recommends that before using the assay in routine TB treatment and control, the performance of the assay in relation to the locally circulating M. tuberculosis bacteria should be validated.

5.6 Novel first-line drug resistance assays

As line-probe assays are somewhat complex to perform and require highly skilled personnel for accurate testing, decentralizing line-probe assays beyond reference laboratories is currently not feasible in most low-income countries. To bring rapid molecular testing for MDR-TB closer to the microscopy level, FIND is partnering with Cepheid on the development of an application for TB and rifampin-resistance detection on its GeneXpert platform [51]. This system is self-contained, fully integrated, and automated platform that combines onboard sample preparation with real-time PCR amplification and detection. The system is designed to purify, concentrate, detect, and identify targeted nucleic acid sequences, delivering results in less than 100 minutes directly from unprocessed samples. Because the system is self-contained, there is no need for a biosafety cabinet and technician training is minimal. In preliminary studies, sensitivity and TB detection was nearly 100% for AFB smear-positive, culture-positive specimens and 70% to 80% for smear-negative, culturepositive specimens, with high test specificity (FIND, unpublished data). Sensitivity and specificity for rifampin resistance has been high. WHO endorsed Xpert MTB/RIF assayin December 2010 and is monitoring the global roll-out of the technology to promote coordination [52].

5.7 Second-line drug resistance assays

The new GenoType Mycobacterium TB drug resistance second-line (MTBDRsl) assay (HainLifescience, Nehren, Germany) provides a potential new tool for the detection of XDR strains of M. tuberculosis within 1–2 days directly in clinical specimen, which could be particularly useful in settings with a high risk of XDR-TB [3]. Diagnosis of latent TB infection specifically caused by drug-resistant strains is currently not possible. Yet, early diagnosis and exact identification of drug resistance during M.tuberculosis latency could have a substantial impact on TB control. The early diagnosis of drug resistance in M. tuberculosis infection should be a priority target of future research activities.

5.8 Diagnosis of latent TB infection

Two interferon-gamma release assays (IGRAs) assays for the diagnosis of LTBI have been commercialized and are undergoing further development to make the application more feasible in field settings. These are the QuantiFERON-TB Gold In-Tube test, a whole blood IGRA from Cellestis, Inc., and the T-Spot.TB test, an Enzyme-linked immunosorbent spot (ELISPOT) assay from Oxford Immunotech, Inc [33]. These assays use the proteins ESAT-6 and CPF-10 from a section of the mycobacterial genome designated RD1 for being a “region of deletion” from Mycobacterium bovis BCG. Such RD1 genes are also absent from most strains of nontuberculous mycobacteria. Thus, the specificity of these tests compared to that of the TST that uses purified protein derivative (PPD) is high, especially in persons with prior BCG vaccination. Study data also suggest that these assays are as or more sensitive than, the TST, especially in immunocompromised persons, including HIV-infected persons. In low-incidence countries where testing for LTBI and treating infected persons at high risk of progression to active TB is an important component of TB control, guidelines on the use of IGRAs, either to supplement or replace the TST, have been issued. It is widely expected that with time IGRAs will replace the TST in screening for LTBI.

 6.   International public databases on Tuberculosis drug resistance mutations

Centralized databases, curated from the literature or high-throughput experiments, have helped user in the era of systems biology, and have helped researchers identify global trends and important biological features that would have only been apparent through a comprehensive analysis of the combined data. Examples include protein–protein interaction databases, genomic databases, and microarray databases.

6.1 TB Drug Resistance Mutation Database (http://www.tbdreamdb.com/)

TBDReaMDB is a comprehensive resource on drug resistance mutations in M. tuberculosis [4]. A systematic review is conducted to identify drug resistance mutations from the existing literature to include in the database. No a priori decisions are made as to whether the mutations described in the literature actually confer drug resistance or are possible secondary compensatory mutations, but instead all mutations that have been described in drug-resistant strains more often than drug-sensitive isolates are included. For each mutation, the database provides complete codon changes for each mutation at both the nucleotide and amino acid level.

The database is divided into two parts. The first part lists all the unique mutations reported in drug-resistant TB isolates, as well as information on the time period of isolate collection, country of origin, molecular detection method, resistance pattern, and susceptibility testing method. Since many of the mutations are reported in multiple publications, only the first report that identified a specific mutation is included.

The second part of the database provides data on the relative frequency of the most common mutations associated with resistance to specific drugs, as reported in surveys from diverse geographical sites. For each drug it contains data from ten high-quality publications that reported the frequency of mutations associated with resistance to that drug. In addition to the information about the frequency of specific mutations the data include time period of isolate collection and country of origin.

Figure 13. The TBDReamTB web site interface.

An interactive Web site allows users to visualize all the specific mutations associated with resistance to each drug. This Web site, in addition to presenting the literature in a manually handled database, also serves as a gateway to post data from future research and development undertaken by its users.

The Web site is organized by drug and presents all the information about the mutations described in the literature. The mutations presented have been color coded to highlight the high-confidence mutations for which frequency data are available. The data are also accessible in the form of global Excel and tab-delimited spreadsheets for all mutations.

Novel mutations found in clinical isolates of drug-resistant TB are included if they (1) are from published studies of clinical M. tuberculosis isolates, (2) occur in isolates that have been characterized by phenotypic drug sensitivity testing, and (3) are identified by specification of the gene, nucleotide position, and the nucleotide and/or amino acid change. Mutations that meet these criteria may be submitted through the Web site and will be added continuously after review.

For studies to be included in the database describing the frequency of common mutations associated with drug resistance in M. tuberculosis, they must (1) be studies of clinical M. tuberculosis isolates; (2) have large sample sizes (a minimum of 100 resistant isolates are required for isoniazid and rifampicin; for the other drugs sample sizes have been determined empirically depending on the ten largest studies published so far—see Web site); (3) report on phenotypic drug sensitivity testing for all isolates; (4) use validated methods to identify drug resistance mutations; (5) identify the nucleotide position and the nucleotide change; and (6) specify the number of resistant and sensitive isolates carrying a specific mutation.

The frequency database is updated regularly with the most recent data.

6.2 The Tuberculosis Database (http://www.tbdb.org/)

The Tuberculosis Database is an integrated database providing access to TB genomic data and resources, relevant to the discovery and development of TB drugs, vaccines and biomarkers [53]. TBDB houses both annotated genome sequence data and microarray and RT–PCR expression data from in vitro experiments and TB-infected tissues. These data and annotations include publicly available sequences from a number of sequencing centers and groups, including sequences being produced by the Broad Institute’s Microbial Sequencing Center. The microarray data within TBDB are predominantly from M. tuberculosis, but we are in the process of incorporating in vivo data from infected host tissues (principally human, primate and murine) into TBDB. Experimental data may be deposited into TBDB by any TB researcher prior to publication providing prepublication access to tools for the analysis, annotation, visualization and sharing of data. The data are then made public at the author’s request or following publication, whichever is first. In addition, TBDB curators search the literature for publications containing relevant TB or host microarray data. The primary data are then requested from the authors of such publications and are entered into TBDB, where the experiments are annotated and made public so other researchers can reanalyze the data (often in conjunction with other datasets within TBDB) using TBDB tools.

References:

  1. World Health Organization. Tuberculosis Fact sheet N°104. – Mode of access: http://www.who.int/mediacentre/factsheets/fs104/en/index.html
  2. World Health Organization. Global tuberculosis control 2011. – Mode of access: http://www.who.int/tb/publications/global_report/en/
  3. Chiang C.-Y., Centis R, Migliori G.B. Drug-resistant tuberculosis: Past, present, future. Respirology (2010) 15, pp. 413–432
  4. Sandgren A [et al]. Tuberculosis Drug Resistance Mutation Database. PLoS Med 6(2) (2009).
  5. Fox W, Ellard GA, Mitchison DA. Studies on the treatment of tuberculosis undertaken by the British Medical Research Council Tuberculosis Units, 1946-1986, with relevant subsequent publications. Int J Tuberc Lung Dis 1999; 3 :S231-79.
  6. Buttger E.C. Drug Resistance in Mycobacterium tuberculosis:Molecular Mechanisms and Laboratory Susceptibility Testing. Donald PR, van Helden PD (eds):Antituberculosis Chemotherapy. Prog Respir Res. Basel, Karger, 2011, vol 40, pp 128–144.
  7. National Institute of Allergy and Infectious Diseases. – Mode of access: http://www.niaid.nih.gov/Pages/default.aspx
  8. Espinal MA, Kim SJ, Suarez PG et al. Standard short-course chemotherapy for drug-resistant tuberculosis. Treatment outcomes in 6 countries. JAMA2000; 283: 2537–45.
  9. Migliori GB, Espinal M, Danilova ID et al. Frequency of recurrence among MDR-TB cases ‘successfully’ treated with standardised short-course chemotherapy. Int. J. Tuberc. Lung Dis. 2002; 6: 858–64.
  10. Quy HTW, Lan NTN, Borgdorff MW et al. Drug resistance among failure and relapse cases of tuberculosis: is the standard re-treatment regimen adequate? Int. J. Tuberc. Lung Dis. 2003; 7: 631–6.
  11. Cole S. Comparative and functional genomics of the Mycobacterium tuberculosis complex. Microbiology. 2002 Oct;148(Pt 10):2919-28.
  12. Cole, S. T., R. Brosch, J. Parkhill, T. Garnier, C. Churcher, et al.1998. Deciphering the biology of Mycobacterium tuberculosis from the complete genome sequence. Nature 393:537-544
  13. MicrobeWiki – Mode of access: http://microbewiki.kenyon.edu/index.php/Mycobacterium
  14. Camus, J.-C., Pryor, M. J., Médigue, C. & Cole, S. T. (2002). Re-annotation of the genome sequence of Mycobacterium tuberculosis H37Rv. Microbiology148, 2967-2973.
  15. Glickman MS, Jacobs WR (February 2001). Microbial pathogenesis of Mycobacterium tuberculosis: dawn of a discipline. Cell 104 (4): 477–85
  16. Kloss P., Xiong L., Shinabarger D.L., Mankin A.S. Resistance mutations in 23 S rRNA identify the site of action of the protein synthesis inhibitor linezolid in the ribosomal peptidyl transferase center. Journal of Molecular Biology. Vol. 294 (1), 19 November 1999, pp. 93–101.
  17. Guide to the Application of Genotyping to Tuberculosis Prevention and Control. – Mode of access:http://www.cdc.gov/tb/programs/genotyping/Chap3/3_CDCLab_2Description.htm
  18. Conell N.D., Kreiswirth B.N. Mycobacterial Strain Genotyping. Tuberculosis A Comprehensive International Approach. Marcel Dekker, 2000
  19. Homolka S, Niemann S, Russell D.G., Rohde K.H. Functional Genetic Diversity among Mycobacterium tuberculosis Complex Clinical Isolates: Delineation of Conserved Core and Lineage-Specific Transcriptomes during Intracellular Survival. PLoS Pathog 6(7) (2010)
  20. Filliol I., Global Phylogeny of Mycobacterium tuberculosis Based on Single Nucleotide Polymorphism (SNP) Analysis: Insights into Tuberculosis Evolution, Phylogenetic Accuracy of Other DNA Fingerprinting Systems, and Recommendations for a Minimal Standard SNP Set. J Bacteriol. 2006 January; 188(2): 759–772.
  21. Brudey K. [et al.] Mycobacterium tuberculosis complex genetic diversity: mining the fourth international spoligotyping database (SpolDB4) for classification, population genetics and epidemiology. BMC Microbiology 2006 6:23
  22. Ramaswamy S, Musser JM (1998) Molecular genetic basis of antimicrobial agent resistance inMycobacterium tuberculosis: 1998 update. Tuber Lung Dis79: 3–29.
  23. Middlebrook G, Cohn ML. Some observations on the pathogenicity of isoniazid-resistant variants of tubercle bacilli. Science 1953;118: 297–9.
  24. Ordway DJ, Sonnenberg MG, Donahue SA et al. Drug-resistant strains of Mycobacterium tuberculosis exhibit a range of virulence for mice. Infect. Immun. 1995; 63: 741–3.
  25. Agerton T., Valway S., Blinkhorn R., et al. Spread of Strain W, a Highly Drug-Resistant Strain of Mycobacterium tuberculosis, Across the United States.Clinical Infectious Diseases, 1999; 29:85-92.
  26. Zhang Y, Yew WW. Mechanisms of drug resistance in Mycobacterium tuberculosis. Int. J. Tuberc. Lung Dis. 2009; 13: 1320–30.
  27. Mendez J. C. Multi Drug Resistance in Tuberculosis and the Use of PCR for Defining Molecular Markers of Resistance. Jacksonville Medicine. 2 (2001).
  28. Valim A., Rossetti M., Ribeiro M, et al. Mutations in the rpoB Gene of Multidrug-Resistant Mycobacterium tuberculosis Isolates from Brazil. J. Clin. Microbiology; Aug. 2000, 3119-3122
  29. Musser J. Antimicrobial Agent Resistance in mycobacteria: molecular genetic insights. Clin Microbiol Rev 1995; 8:496-514.
  30. Telenti A., Honore N., Bernasconi C., et al. Genotypic assessment of isoniazid and rifampin resistance in Mycobacterium tuberculosis: a blind study at reference laboratory level. J. Clin Microbiol 1997; 35:719-723
  31. Huyen M.N. [et al.] Validation of the GenoType MTBDRplus assay for diagnosis of multidrug resistant tuberculosis in South Vietnam. BMC Infectious Diseases 2010, 10:149
  32. Musser J.M., Kapur V., Williams D.L., Kreiswirth B.N., Soolingen D., Embden J. Characterization of the Catalase-Peroxidase Gene (katG) and inhA Locus in Isoniazid-Resistant and -Susceptible Strains of Mycobacterium tuberculosis by Automated DNA Sequencing: Restricted Array of Mutations Associated with Drug Resistance. The Journal ofInfectious Diseases 1996;173:196-202
  33. Seung K., Rich M.L. Diagnosis and Treatment of Drug-Resistant Tuberculosis. Tuberculosis: the essentials, Fourth edition, 2009.
  34. Davies AP, Billington OJ, McHugh DA, et al. Comparison of Phenotypic and Genotypic Methods for Pyrazinamide Susceptibility Testing with Mycobacterium tuberculosis. J Clin Microbiol 2000; 38(10):3686–3688.
  35. World Health Organization. Guidelines of Drug Susceptibility Testing for Second-Line Anti-Tuberculosis Drugs in DOTS-Plus Programs; (WHO/CDS/TB/2001.288). Geneva, Switzerland: World Health Organization, 2001.
  36. American Thoracic Society/Centers for Disease Control and Prevention/Infectious Diseases Society of America. Treatment of tuberculosis. Am J Respir Crit Care Med 2003; 167(4): 603–662.
  37. Updated Guidelines for the Use of Nucleic Acid Amplification Tests in the Diagnosis of Tuberculosis. – Mode of access:http://www.cdc.gov/mmwr/preview/mmwrhtml/mm5801a3.htm
  38. Boskey E. Commercial Nucleic-Acid Amplification Tests for Diagnosis of Pulmonary Tuberculosis in Respiratory Specimens: Meta-Analysis and Meta-Regression. – Mode of access: http://std.about.com/od/M-O/g/Nucleic-Acid-Amplification-Tests.htm
  39. World Health Organization. Guidelines for the Programmatic Management of Drug-Resistant Tuberculosis, Emergency Update. Geneva, Switzerland:World Health Organization, 2008
  40. Zhang Y, Yew WW. Mechanisms of drug resistance in Mycobacterium tuberculosis. Int. J. Tuberc. Lung Dis. 2009; 13: 1320–30.
  41. Barnard M, Albert H, Coetzee G et al. Rapid molecular screening for multidrug-resistant tuberculosis in a high-volume public health laboratory in South Africa. Am. J. Respir. Crit. Care Med. 2008; 177: 787–92.
  42. Nolte FS, Metchock B, McGowan JE et al. Direct detection of Mycobacterium tuberculosis in sputum by polymerase chain reaction and DNA hybridization. J. Clin. Microbiol. 1993; 31: 1777–82.
  43. Lin SYG, Probert W, Lo M et al. Rapid detection of isoniazid and rifampin resistance mutations in Mycobacterium tuberculosis complex from cultures or smear-positive sputa by use of molecular beacons. J. Clin. Microbiol. 2004; 42: 4204–8.
  44. Caviedes L, Lee TS, Gilman RH et al. Rapid, efficient detection and drug susceptibility testing of Mycobacterium tuberculosis in sputum by microscopic observation of broth cultures. J. Clin. Microbiol. 2000; 38: 1203–8.
  45. Hamid Salim A, Aung KJM, Hossain MA et al. Early and rapid microscopy-based diagnosis of true treatment failure and MDR-TB. Int. J. Tuberc. Lung Dis. 2006; 10:1248–54.
  46. Pai M, Kalantri S, Pascopella L et al. Bacteriophage-based assays for the rapid detection of rifampicin resistance in Mycobacterium tuberculosis: a meta-analysis. J. Infect. 2005; 51: 175–87.
  47. Martin A, Portaels F, Palomino JC. Colorimetric redox-indicator methods for the rapid detection of multidrug resistance in Mycobacterium tuberculosis: a systematic review and meta-analysis. J. Antimicrob. Chemother. 2007; 59: 175–83.
  48. Martin A, Cubillos-Ruiz A, Von Groll A et al. Nitrate reductase assay for the rapid detection of pyrazinamide resistance in Mycobacterium tuberculosis using nicotinamide. J. Antimicrob. Chemother. 2008; 61: 123–7.
  49. Piatek A.,Telenti A.,Murray M., et al. Genotypic Analysis of Mycobacterium tuberculosis in Two Distinct Populations Using Molecular Beacons: Implications for Rapid Susceptibility Testing. Antimicrobial Agents and Chemotherapy, Jan 2000; 44(1): 103-110.
  50. Molecular line probe assays for rapid screening of patients at risk of multi-drug resistant tuberculosis (MDR-TB). World Health Organization, 2008.
  51. GeneXpert System. Molecular diagnostics made fast, accurate and easy. – Mode of access: http://www.cepheid.com/systems-and-software/genexpert-system
  52. Rapid Implementation of Xpert MTB/RIF. WHO, Geneva WHO/HTM/TB/2011.2, 2011. – Mode of access:http://whqlibdoc.who.int/publications/2011/9789241501569_eng.pdf
  53. Reddy TB [et al.] TB database: an integrated platform for tuberculosis research. Nucleic Acids Res (2009) 37.