Tuesday, November 28, 2006
Typhoid Bacteria - Resistant Strains
Date:November 28, 2006
In a study published in the latest issue of Science (24 November, 2006), an international consortium from the Max-Planck Society, Wellcome Trust Institutes in Britain and Vietnam, and the Institut Pasteur in France have elucidated the evolutionary history of Salmonella Typhi.
Typhi is the cause of typhoid fever, a disease that sickens 21 million people and kills 200,000 worldwide every year. The results indicate that asymptomatic carriers played an essential role in the evolution and global transmission of Typhi. The rediscovered importance of the carrier state predicts that treatment of acute disease, including vaccination, will not suffice to eradicate this malady. The results also illuminate patterns leading to antibiotic resistance after the indiscriminate use of antibiotics. Fluoroquinolone treatment in southern Asia over two decades has resulted in the emergence of multiple, independent nalidixic acid-resistant mutants, of which one group, H58, has multiplied dramatically and spread globally. The prevalence of these bacteria hampers medical cure of clinical disease via antibiotics.
Salmonella Typhi. (Image: Dr. Volker Brinkmann, MPI for Infection Biology)
Typhoid fever remains a major health problem in the developing world and continues to cause disease in Europe and on the american continent. The evolutionary history and population structure of Typhi were poorly understood, partly because these bacteria show little genetic diversity. Now a team led by Mark Achtman and Philippe Roumagnac from the Max Planck Institute for Infection Biology, Berlin, has applied population genetic experience from prior work with Yersinia pestis, Escherichia coli, Helicobacter pylori and Neisseria meningitidis to provide novel insights into the evolution of this pathogen.
The team combined its resources to assemble for the first time a globally representative collection of 105 strains of Typhi and investigated the sequence diversity within 90,000 base pairs per strain. Eighty-eight informative sequence differences were detected, showing that the population structure has evolved over the last 10,000 to 43,000 years. Amazingly, the ancestral strain continues to exist today, as do many of its direct descendents, indicating a neutral population structure, whereas normally selective forces lead to extinction of intermediate genotypes. Furthermore, these bacteria are distributed globally, demonstrating that Typhi has spread inter-continentally on multiple occasions.
The authors propose that the unusual population structure of Typhi reflects long-term carriage by asymptomatic carriers, who reached public notoriety at the beginning of the 20th century with "Mr. N the milker" in England and Typhoid Mary (Mary Mallon) in the U.S.A. These individuals infected 100s of people over the decades while they worked in the food production industry. Healthy carriers may have allowed Typhi to survive in hunter-gatherer populations prior to the Neolithic expansion of city states and facilitated its intercontinental spread. Healthy carriers are also consistent with the observation that individual genotypes of Typhi persist for many decades within each country.
Increasing resistance to antibiotics in recent decades has hampered efforts of clinicians to cure typhoid fever. The indiscriminate use of fluoroquinolones, which is a cost-effective, standard treatment for typhoid fever, has been accompanied by a frightening increase in the numbers of resistant Typhi. Investigations of a large strain collection from southern Asia revealed that many different genotypes independently acquired resistance to nalidixic acid, a quinolone. One of these genotypes, H58, has become predominant throughout southern Asia and has even spread to Africa. In Vietnam, up to 95% of Typhi are now resistant to nalidixic acid and many other antibiotics. Although these cases can still be treated with newer antibiotics, those antibiotics are much more expensive than standard fluoroquinolones, which raises the cost of medical treatment. Furthermore, it is likely that Typhi will develop resistance to these antibiotics as well.
The combination of these investigations raises problems for public health measures. Indiscriminate antibiotic usage results in real-time evolution of bacteria that resist treatment. Furthermore, the healthy carrier state provides a safe reservoir for these bacteria which allows them to evade short-term antibiotic treatment and vaccination, indicating that typhoid fever will remain a major health problem for the foreseeable future.
The research was carried out collaboratively by the Max Planck Institute for Infection Biology in Berlin, the Wellcome Trust Sanger Institute, Hinxton Hall, the Institut Pasteur, Paris, and the Oxford University Clinical Research Unit, Ho Chi Minh City, with assistance from hospitals in Ho Chi Minh City and Hanoi and the International Vaccine Institute in Seoul. Financial support was by the Wellcome Trust, UK.
Tuesday, November 07, 2006
Development and evaluation of a real-time PCR assay targeting the type III secretion system of Burkholderia pseudomallei.
1: J Clin Microbiol. 2006 Jan;44(1):85-90.
Ryan T. Novak,1 Mindy B. Glass,1* Jay E. Gee,1 Daniel Gal,2 Mark J. Mayo,2 Bart J. Currie,2 and Patricia P. Wilkins1
Meningitis and Special Pathogens Branch, Division of Bacterial and Mycotic Diseases, National Center for Infectious Diseases, Centers for Disease Control and Prevention, Atlanta, Georgia 30333,1 Menzies School of Health Research, Charles Darwin University, Northern Territory Clinical School, Flinders University, Royal Darwin Hospital, Darwin, Northern Territory, Australia2
Received 21 June 2005/ Returned for modification 19 July 2005/ Accepted 11 October 2005
Here we report on the development of a discriminatory real-time assay for the rapid identification of Burkholderia pseudomallei isolates and the evaluation of this assay for sensitivity against related species and detection in spiked human blood samples. The assay targets a 115-base-pair region within orf2 of the B. pseudomallei type III secretion system gene cluster and distinguishes B. pseudomallei from other microbial species. Assay performance was evaluated with 224 geographically, temporally, and clinically diverse B. pseudomallei isolates from the Centers for Disease Control and Prevention strain collection. This represents the first real-time PCR for rapid and sensitive identification of B. pseudomallei that has been tested for cross-reactivity with 23 Burkholderia mallei, 5 Burkholderia thailandensis, and 35 Burkholderia and 76 non-Burkholderia organisms which have historically presented diagnostic challenges. The assay performed with 100% specificity. The limit of detection was found to be 76 femtograms of DNA (equivalent to 5.2 x 103 genome equivalents per ml) in a single PCR. In spiked human blood, the assay could detect as few as 8.4 x 103 CFU per ml. This rapid assay is a valuable tool for identification of B. pseudomallei and may improve diagnosis in regions endemic for melioidosis.
Melioidosis, a disease caused by infection with Burkholderia pseudomallei, is a significant public health burden in Southeast Asia and northern Australia, where in some regions it is the most common cause of fatal community-acquired bacteremic pneumonia (6, 45). The ecological distribution of B. pseudomallei is mostly limited to the tropical and subtropical areas between 20° north and south latitudes. While occasional cases of melioidosis occur beyond these regions, outside the tropics they have usually been associated with imported animals or have occurred in travelers returning from these regions (6, 7, 12). Clinical manifestations of melioidosis range from asymptomatic infection to focal infections in skin, muscle, bones, joints, or internal organs to pneumonia, which can be chronic or fulminant with rapidly fatal septicemia, where death can occur within 48 h of symptom onset (6, 8, 45). B. pseudomallei infection can resemble a number of other infections including typhoid, tuberculosis, and malaria (45). High infectivity, severity of disease, and environmental persistence have resulted in B. pseudomallei being classified as a category B select agent by the Centers for Disease Control and Prevention (CDC) (31).
Culture remains the current "gold standard" for diagnosis of melioidosis and identification of B. pseudomallei from a clinical specimen (43, 48, 50). However, definitive identification relies on a battery of traditional microbiological tests that can require up to a week to complete. Furthermore, some biochemical identification methods for culture isolates are problematic, especially rapid biochemical assays which have been shown to misidentify B. pseudomallei as other bacteria such as Pseudomonas spp., Burkholderia vietnamiensis, Stenotrophomonas maltophilia, and in particular Chromobacterium violaceum, which can cause an infection with similar symptoms to melioidosis (14, 19, 23). Available serologic assays (dot immunoassay, indirect hemagglutination assay, enzyme-linked immunosorbent assay, and immunochromatograpic test) are not particularly useful in regions endemic for melioidosis because past exposure to B. pseudomallei may result in seroconversion (6, 27, 45). Rapid bacterial detection from clinical samples using latex agglutination or direct immunofluorescence has been successful in Thailand, but these assays are not commercially available (42, 49). Compounding the need for rapid diagnosis is the lack of a human vaccine and the intrinsic resistance of B. pseudomallei to many of the affordable antibiotic treatments for general sepsis in areas endemic for the disease (7, 20, 45). In addition to accelerating appropriate therapeutic intervention, improved rapid diagnostic tests could potentially prevent unnecessary laboratory-acquired infection due to overhandling of clinical specimens containing B. pseudomallei and would facilitate surveillance prior to and following deliberate release (3, 5, 7, 31).
The complete sequence of the B. pseudomallei genome reveals several gene targets for the identification of this species (e.g., 16S rRNA genes, fliC, rpsU, and type III secretion [TTS] gene cluster) (18, 25, 26, 36, 38, 39). This approach to PCR target identification and primer and probe design and improved methods for fluorescence labeling of DNA allow for the in silico design and evaluation of real-time PCR assays specific for a given species (15, 24, 29, 30). The disadvantage of a purely theoretical approach is choosing gene targets based on one strain, which may not be representative of the global B. pseudomallei population, resulting in false-negative results. Thus for this study we chose a target based on not only its theoretical specificity but also its apparent ubiquitous population distribution (35). Previously, a 548-bp region of the B. pseudomallei type III secretion system (TTS1) gene cluster encompassing part of open reading frame 2 (orf2) was found to be present in B. pseudomallei and not in the related Burkholderia mallei or Burkholderia thailandensis (28, 46, 47). This open reading frame was later found to be present in all of a large number of northern Australian B. pseudomallei strains (35). We report here the design and evaluation of a fluorescence-labeled-probe-based real-time PCR assay targeting orf2 of the TTS1 of B. pseudomallei (TTS1 real-time PCR). The sensitivity, specificity, low limit of detection, and lack of interaction with human DNA make this assay a good candidate for diagnostic clinical laboratory evaluation.
Materials and Methods
Collection and preparation of bacterial strains.
Bacterial strains were obtained from CDC culture collections housed in the Meningitis and Special Pathogens Branch. They included 224 B. pseudomallei strains representing temporal (1949 to 2004), geographic (Americas, Asia, Europe, Africa, and Oceania), and origin diversity (human, environmental, animal). The geographic regions covered by this collection represent the locale of isolation and not necessary where infection occurred. Isolates from regions not traditionally considered endemic for melioidosis such as North America and Europe represent imported cases. In addition, 23 B. mallei strains, 5 B. thailandensis strains, and 111 non-B. pseudomallei strains were evaluated for cross-reactivity (Table 1). All strains were stored at –70°C in defibrinated rabbit blood until they were tested. Identification of strains was carried out with a standard battery of biochemical tests (44). Culture isolates were not available from six of the diagnostically challenging organisms (Mycobacterium tuberculosis, Francisella tularensis, Plasmodium falciparum, Plasmodium vivax, Plasmodium ovale, and Plasmodium malariae). For these pathogens DNA was obtained from subject matter expert laboratories and tested directly.
Isolates were cultured on standard media at 37°C overnight. For testing assay specificity whole-cell DNA extractions were prepared from the cultures by heat lysis as previously described by Hoffmaster et al. (17). The assay sensitivity was evaluated using purified B. pseudomallei genomic DNA extracted with the DNeasy tissue kit (QIAGEN, Valencia, CA) using the manufacturer's instructions for gram-negative bacteria. For spiked human blood samples, DNA was isolated from a 200-µl sample using a modified protocol for the QIAmp DNA blood kit (QIAGEN), substituting proteinase K for protease and buffer ATL for AL and using an elution volume of 200 µl. The procedure was modified to add a 10-min 70°C step to ensure lysis of bacterial cells. The DNA concentrations were determined using a spectrophotometer (NanoDrop Technologies, Wilmington, DE), and the numbers of genome equivalents (GE) were calculated using the 7.25-Mbp genome (68% G+C) of B. pseudomallei K96243 (18).
Real-time PCR assay. Primer pair and probe were designed to target the type III secretion system gene cluster of B. pseudomallei (GenBank accession no. AF074878) using Primer Express 2.0 (Applied Biosystems, Inc., Foster City, CA), BpTT4176F (5'-CGTCTCTATACTGTCGAGCAATCG-3'), and BpTT4290R (5'-CGTGCACACCGGTCAGTATC-3'). The primer pair was designed to generate a 115-bp PCR product. The fluorogenic probe BpTT4208P (5'-CCGGAATCTGGATCACCACCACTTTCC-3') was synthesized with a 6-carboxyfluorescein reporter molecule attached at the 5' end and a Black Hole Quencher 1 on the 3' end. The primers and probe were synthesized in the Biotechnology Core Facility at the CDC.
Each TTS1 real-time PCR mixture contained a 2-µl aliquot of extracted DNA in 23 µl of a PCR master mixture containing 1x LightCycler Faststart DNA master hybridization probe reaction mixture (Roche Applied Sciences, Indianapolis, IN), 5 mM MgCl2, 400 nM of each primer, and 200 nM of probe. Amplification and detection were performed with a SmartCycler (Cepheid, Sunnyvale, CA) using the standard settings. The SmartCycler thermal cycling conditions were 95°C for 8 min, followed by 45 cycles of 15 s at 95°C and 15 s at 59°C. The cycle threshold values represent the calculated cycles at which fluorescence from the cleaved probe exceeds a fixed threshold, as calculated by the instrument for each reaction. No-template (PCR grade water) and positive (B. pseudomallei CDC2000032028) controls were included in each run to rule out the possibility of contamination or amplification failure. The sensitivity and specificity of the real-time PCR assay were evaluated by testing 1 ng of extracted DNA.
The lower limit of detection (LLOD), linear dynamic range (LDR), and assay variance for the TTS1 real-time PCR were determined using purified B. pseudomallei (CDC2000032028) DNA (13). For LDR and variance analysis, five replicates from a 10-fold serial dilution (108 to 103 GE/ml) of purified DNA in elution buffer (QIAGEN, Valencia, CA) were assayed in a single run. To determine LLOD, an additional five replicates from a 10-fold serial dilution (102 to 10–2 GE/ml) of purified DNA were added. Eight separate replicate experiments were conducted using aliquots of DNA frozen at –20°C. These data were compared to those obtained from DNA extractions of six spiked human blood samples (Lampire Biological, Coopersburg, PA, and CDC Blood Services Group, Atlanta, GA). Previous studies have highlighted the need for preassay enrichment of clinical samples to achieve DNA concentrations in the range of real-time detection (38, 39). Therefore, we designed an evaluation of the assay for detecting B. pseudomallei cells in human blood. An aliquot of the uninoculated whole human blood (with EDTA as a non-PCR-inhibiting anticoagulant) was plated prior to each experiment to check for sterility. Colonies from overnight cultures of B. pseudomallei (CDC2000032028) were suspended in phosphate-buffered saline (PBS) at a concentration equivalent to 109 cells per ml based on McFarland turbidimetric standards. Serial 10-fold dilutions (108 to 10–2 CFU/ml) were made from the solution in blood and PBS. Aliquots from each series of dilutions were used for spread plates in triplicate, and the average CFU/ml was calculated. Four replicates of each series were assayed.
Data and statistical analysis.
The LLODs were determined using a model of nonlinear regression (PriProbit version 1.63). The LDR and the coefficient-of-variation calculations for the assay were carried out with Excel (Microsoft). The PCR efficiency (E) of the primer pair and probe was calculated using the equation E = 10–1/slope – 1 (40). An E of 1.0 indicates that amplicon quantity is duplicated every cycle.
Sensitivity and specificity.
The assay successfully amplified DNA from all 224 B. pseudomallei strains in the CDC culture collection. The cycle threshold values ranged from 19 to 27 for the whole-cell extracts. Using a nonlinear-regression model, the LLOD from purified B. pseudomallei (CDC2000032028) DNA was 76 fg/PCR (equivalent to 5.2 x 103 GE/ml) (Fig. 1A), and the LDR was 3.3 x 104 GE/ml to 3.3 x 108 GE/ml (Fig. 1B). Intra-assay variance was estimated in the LDR to be between 0.39 and 1.64%, while the interassay variability was between 1.00 and 1.76%.
The specificity of the designed primers and probe was assessed first in silico by comparing the sequences and the gene product using BLAST (2). There were no sequences in the database producing alignments with significant expectation values or sharing regions of significant similarity to the primers or probe.
All non-B. pseudomallei isolates were negative (Table 1). These included genetic relatives (e.g., Burkholderia mallei, B. thailandensis, and Ralstonia spp.), biothreat agents (e.g., Bacillus anthracis, Brucella spp., and Francisella tularensis), and pathogens that are part of the differential diagnoses for melioidosis (e.g., Staphylococcus aureus, Plasmodium spp., Salmonella spp., and Mycobacterium tuberculosis). Organisms that cause frequent misidentifications using rapid biochemical assays (e.g., Pseudomonas spp., Burkholderia vietnamiensis, Burkholderia cepacia, Stenotrophomonas maltophilia, and Chromobacterium violaceum) were negative by the TTS1 real-time assay.
Assay of spiked blood samples.
Human DNA was nonreactive by the TTS1 real-time PCR. Five separate experiments were performed using spiked blood samples from five different donors with heterogeneous blood types and travel histories. Mean assay results were used to establish an LLOD of 8.4 x 103 CFU/ml (Fig. 2A) and the LDR of 5.4 x 104 to 5.4 x 108 CFU/ml (Fig. 2B). The regression coefficient (R2 = 0.99) and the amplification efficiency value (E = 1.17) demonstrated an exponential amplification of DNA with this probe and primer pair.
Melioidosis is a significant public health burden in regions of Southeast Asia and northern Australia. Septicemic melioidosis, if left untreated, is a rapidly progressing infection that has a mortality rate as high as 90% (45). In regions not endemic for melioidosis Burkholderia pseudomallei is also of concern because of its potential release during a biothreat event. Therefore, we developed a real-time PCR for rapid identification of B. pseudomallei isolates that potentially offers a faster and more reliable diagnosis of disease in regions endemic for melioidosis. Rapid identification of B. pseudomallei is especially important in these regions because of the high prevalence of B. pseudomallei strains that are resistant to the antibiotics typically used to empirically treat sepsis (20, 45). Other methods of rapid identification such as latex agglutination are not commercially available. Here we have described the development and evaluation of a rapid, sensitive real-time assay for the specific identification of B. pseudomallei. The real-time assay targeting orf2 of the B. pseudomallei type III secretion system accurately distinguished a large sample of diverse B. pseudomallei isolates from non-B. pseudomallei isolates (100% specificity) and was not inhibited by human blood products or DNA.
A number of rapid presumptive biochemical and PCR assays have been developed and evaluated for the identification of B. pseudomallei (14, 16, 21, 27, 32, 51). The molecular methods for identification of B. pseudomallei infection include a number of conventional PCRs with various specificities that also may suffer from a lack of sensitivity (16, 22, 32, 51). More recently two real-time PCR assays have been developed and well evaluated on bacterial isolates (36, 38, 39). Both real-time assays had greater sensitivity than the traditional PCR assays and appear specific for the identification of isolates of B. pseudomallei or the B. pseudomallei/B. mallei complex. This increased sensitivity suggests that they may be suitable for clinical samples, but both assays may require DNA or cell preassay enrichment to detect bacteremic infections. There have been no reported evaluations of either assay for diagnosis of melioidosis in a clinical setting.
Estimates of the concentration of B. pseudomallei in blood for septicemic melioidosis patients range from 1 to 1,000 CFU per ml (33, 37, 41). The onset of melioidosis symptoms for a given blood cell count varies depending on a combination of isolate virulence and host factors (9, 10, 35). Based on experiments with B. pseudomallei-spiked blood the TTS1 real-time assay has a 95% probability of detecting a concentration of 8.4 x 103 CFU/ml with a single PCR (Fig. 2A). Following analysis of DNA extracted from multiple dilutions of B. pseudomallei-spiked human blood samples, the lowest concentration that yielded at least one positive target amplification was 500 CFU/ml, equaling a 12.5% probability of detection (Fig. 2A). By increasing the assay volume of extracted clinical DNA in this system fivefold and testing multiple replicates, the probability of detecting a positive clinical sample using the TTS1 real-time assay is likely to improve. Assuming a Poisson distribution of sample DNA, a clinical specimen with 500 cells/ml has an extrapolated probability of detection of 100% in at least two of three replicate samples with this fivefold increase in sample volume.
These results suggest that early bacteremia may be beyond the limit of detection of the current assay conditions. Data presented here were based on DNA extracted from spiked blood samples; no attempt to concentrate the cells or DNA was made prior to extraction. Early detection may be accomplished by assaying a concentrated sample either through preprocessing of a blood specimen or collecting a clinical specimen with a higher cell concentration such as sputum or wound exudate from a focal infection site (B. Currie, personal communication). An adjusted assay reduces the time for presumptive diagnosis from days to 3 h, allowing for early definitive antibiotic intervention and minimizing laboratorian contact time.
The power of a well-designed and evaluated real-time PCR assay is demonstrated by its ability to detect target DNA in the presence of a vast excess of other nontarget DNA (24). During outbreaks of melioidosis associated with severe weather events such as cyclones, after natural disasters such as tsunamis, or following a bioterror release, large numbers of patients could potentially overwhelm the capacity of a health care facility (1, 10). A reliable single target assay could rapidly screen large numbers of patients using multiple specimens from usually sterile (blood or urine), and nonsterile (wound, throat, or sputum) clinical sites to identify those needing immediate antibiotic therapy for melioidosis (34).
Testing environmental samples containing diverse microbial ecology may be enhanced by the use of a selective identification technique such as the TTS1 real-time PCR, but this requires further evaluation. Also, large-scale environmental surveys to establish ecological distribution and concentration using this assay would be a benefit in endemic areas for melioidosis.
The real-time PCR assay targeting orf2 of the B. pseudomallei type III secretion system is a powerful tool for the rapid identification of B. pseudomallei and can potentially complement current confirmatory diagnostic procedures for melioidosis. Further studies to evaluate the TTS1 real-time assay on a larger panel of environmental samples, including an evaluation of more B. thailandensis strains, are needed. This less-virulent close relative of B. pseudomallei is a common soil saprophyte in the hyperendemic melioidosis region of northeast Thailand (4, 11). The evaluation of rapid diagnostic assays using clinical samples in a region of melioidosis endemicity is necessary for an accurate assessment of the clinical efficacy of proposed new diagnostic tools and is under way as part of an ongoing prospective melioidosis cohort study in northern Australia.
We thank Glenn Morlock for supplying the Mycobacterium tuberculosis DNA, Alexandre Dasilva for supplying the Plasmodium sp. DNA, and Timothy Barrett for supplying the Salmonella enterica serovar Urbana and Vibrio cholerae strains used in this study.
This research was supported in part by an appointment to the Emerging Infectious Diseases Fellowship Program administered by the Association of Public Health Laboratories and funded by the Centers for Disease Control and Prevention and by an Australian National Health and Medical Research Council Project Grant.
* Corresponding author. Mailing address: Meningitis and Special Pathogens Branch, Division of Bacterial and Mycotic Diseases, National Center for Infectious Diseases, CDC, MS-G34, 1600 Clifton Rd., N.E., Atlanta, GA 30333. Phone: (404) 639-4055. Fax: (404) 639-3023. E-mail: firstname.lastname@example.org
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