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Due to the occurrence of natural plague outbreaks and its historical usage as a biological weapon, Yersinia pestis is considered one of the high-priority biological threat agents. It can remain viable in certain environments including water for >100 days. Because of its slow-growth characteristic, it usually takes three or more days to detect and confirm the identity of viable Y. pestis cells by PCR, serological, or biochemical assays when using the traditional microbiological plate-culture-based analysis, and that too, assuming faster growing microbes present in a water sample do not mask the Y. pestis colonies and interfere with analysis. Therefore, a rapid-viability Polymerase Chain Reaction (RV-PCR) method was developed for detection of Y. pestis. The RV-PCR method combines 24 h-incubation broth culture in a 48-well plate, and pre- and post-incubation differential PCR analyses, thereby allowing for rapid and high-throughput sample analysis compared with the current plate culture method. One chromosomal and two plasmid gene target-based real-time PCR assays were down-selected, showing ca. 10 genome equivalent detection; the chromosomal assay was then used for RV-PCR method development. A 101-cell level (10–99 cells) sensitivity of detection was demonstrated even with complex sample backgrounds including known PCR inhibitors (ferrous sulfate and humic acid), as well as metal oxides and microbes present in Arizona Test Dust (ATD). The method sensitivity was maintained in the presence of dead Y. pestis cells up to 104 cells per sample. While affording high-throughput and rapid sample analysis, the 48-well plate format used in this method for sample enrichment significantly reduced labor requirements and generation of BioSafety Level-3 (BSL-3) laboratory waste as compared to the usual microbiological plate-culture-based methods. This method may serve as a model for other vegetative bacterial pathogens.

Francisella tularensis, which causes potentially fatal tularemia, has been considered an attractive agent of bioterrorism and biological warfare due to its low infectious dose, reported environmental persistence, and ability to be transmitted to humans via multiple exposure routes. Due to slow growth on even selective culture media, detection of viable F. tularensis from environmental and drinking water samples, usually takes > 3 days. Therefore, a rapid viability polymerase chain reaction (RV-PCR) method was developed to detect and identify viable F. tularensis cells in environmental samples. The method uses a change in PCR response during high throughput (48-well) sample incubation in Brain Heart Infusion/Vitox/Fildes/Histidine growth medium to detect viable F. tularensis presence, which is at least two times faster than the current plate culture-based method. Using the method, 10(1) to 10(2) live F. tularensis cells were detected in simulated complex sample matrices containing chemical and biological interferences.

To rapidly remediate facilities after a biothreat agent release, improved turnaround times are needed for sample analysis. Current methods to confirm the presence of a viable biothreat agent are limited by low sample throughput. We have developed a rapid-viability-polymerase chain reaction (RV-PCR) method to determine the presence of viable spores. The method combines high-throughput sample processing with 96-well PCR analysis, which measures a change in real-time, quantitative PCR response arising from increased target-cell populations during culturing. The method accurately detects 1 to 10 live spores in a high-dead spore background (10(6)). Field tests using approximately 1000 biological indicators, each containing 10(6) spores of the B. anthracis surrogate, Bacillus atrophaeus, exposed to seven lethal and sub-lethal chlorine dioxide levels showed no significant difference (p0.05) between RV-PCR and standard culturing methods for detecting the percent survival of spores. RV-PCR results were obtained in17 h compared to 7 days for the standard culturing method. High-throughput sample processing and RV-PCR protocols were also developed and tested for synthetic wipe samples containing reference dirt material. RV-PCR protocols allowed processing and accurate analysis of approximately100 dirty wipe samples (2''x2'' synthetic) containing approximately10 viable B. atrophaeus spores in24 h. Quantitative RV-PCR protocols based on a Most-Probable-Number (MPN) statistical approach developed for B. anthracis Sterne resulted in more rapid turnaround times than those for traditional culturing and no significant difference in log colony-forming units compared to traditional viability analysis. Integration of RV-PCR assays with high-throughput protocols will allow the processing of 200 wipe samples per day per robot using commercially available automation.

A comparison of Most-Probable-Number Rapid Viability (MPN RV) PCR and traditional culture methods for the quantification of Bacillus anthracis Sterne spores in macrofoam swabs from a multi-center validation study was performed. The purpose of the study was to compare environmental swab processing methods for recovery, detection, and quantification of viable B. anthracis spores from surfaces. Results show that spore numbers provided by the MPN RV-PCR method were typically within 1-log of the values from a plate count method for all three levels of spores tested (3.1x10(4), 400, and 40 spores sampled from surfaces with swabs) even in the presence of debris. The MPN method tended to overestimate the expected result, especially at lower spore levels. Blind negative samples were correctly identified using both methods showing a lack of cross contamination. In addition to detecting low levels of spores in environmental conditions, the MPN RV-PCR method is specific, and compatible with automated high-throughput sample processing and analysis protocols, enhancing its utility for characterization and clearance following a biothreat agent release.