Highly Sensitive Direct Detection and Quantification of Burkholderia pseudomallei Bacteria in Environmental Soil Samples by Using Real-Time PCR
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Highly Sensitive Direct Detection and Quantification of Burkholderia pseudomallei Bacteria in Environmental Soil Samples by Using Real-Time PCR
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Authors:
Address: 1Friedrich Loeffler Institute of Medical Microbiology, Ernst Moritz Arndt University of Greifswald, Greifswald, Germany
Journal: Appl Environ Microbiol. 2011 September; 77 (18) : 6486–6494.
Publication:
The Soil bacterium and potential biothreat agent Burkholderia pseudomallei causes the infectious disease melioidosis, which is naturally acquired through Environmental contact with the bacterium. Environmental Detection of B. pseudomallei represents the basis for the development of a geographical risk map for humans and livestock. The aim of the present study was to develop a Highly Sensitive, culture-independent, DNA-based method that allows Direct Quantification of B. pseudomallei from soil. We established a protocol for B. pseudomallei soil DNA isolation, purification, and quantification by quantitative PCR (qPCR) targeting a type three secretion system 1 single-copy gene. This assay was validated Using 40 soil Samples from Northeast Thailand that underwent parallel bacteriological culture. All 26 samples that were B. pseudomallei positive by direct culture were B. pseudomallei qPCR positive, with a median of 1.84 × 104 genome equivalents (range, 3.65 × 102 to 7.85 × 105) per gram of soil, assuming complete recovery of DNA. This was 10.6-fold (geometric mean; range, 1.1- to 151.3-fold) higher than the Bacterial count defined by direct culture. Moreover, the qPCR detected B. pseudomallei in seven samples (median, 36.9 genome equivalents per g of soil; range, 9.4 to 47.3) which were negative by direct culture. These seven positive results were reproduced using a nested PCR targeting a second, independent B. pseudomallei-specific sequence. Two samples were direct culture and qPCR negative but nested PCR positive. Five samples were negative by both PCR methods and culture. In conclusion, our PCR-based system provides a highly specific and sensitive tool for the quantitative environmental surveillance of B. pseudomallei.
The Gram-negative betaproteobacterium Burkholderia pseudomallei, a natural inhabitant of soil and surface water, causes the infectious disease melioidosis, with high mortality rates for infected humans in tropical and subtropical regions where the disease is endemic (7, 10, 58). Clinical cases of melioidosis have been reported regularly in Southeast Asian countries, such as Thailand (28, 53), Vietnam (42), and Malaysia (43, 62), in northern and Western Australia (12, 19), and sporadically in India (1, 49), South and Central America (20, 45), and West and East Africa (21). The disease can be acquired by inoculation of B. pseudomallei through skin lesions, by aerosols from contaminated soil and surface water, or by ingestion (11, 15). Melioidosis has attracted interest outside its known distribution areas by cases in travelers (9) and soldiers (32, 40) returning from regions of B. pseudomallei endemicity and by its classification as a category B bioterrorism agent (47). In many known areas and countries of endemicity where melioidosis is likely to occur, diagnostic resources in the clinical laboratory are limited, and therefore, the true burden of the disease and the worldwide distribution of B. pseudomallei remain unclear.
Apart from the detection of B. pseudomallei in clinical cases, knowledge of the distribution and lifestyle of B. pseudomallei in its natural soil environment is important for understanding the epidemiology of melioidosis. In this context, quantitative detection of B. pseudomallei is crucial for investigating its associations with specific habitats, as well as the influence of factors such as climate change. Several investigations have used culture-based quantification (39, 50) to enumerate B. pseudomallei bacteria within its natural environment. Quantitative cultivation of B. pseudomallei from soil samples depends on efficient detachment of microorganisms from the soil matrix, which relies on the selected dispersion method (56). However, detection of B. pseudomallei by culture can be hindered by the presence and overgrowth of other environmental bacterial species capable of growing on the currently used selective media (6), especially when only low B. pseudomallei cell numbers are present. Additionally, the proportion of B. pseudomallei environmental cells which might be viable but are in a nonculturable state under standard laboratory conditions is unknown. It seems likely that this phenomenon, described for other environmental species (2, 36), also contributes to an underestimation of the B. pseudomallei bacterial load or to false-negative results from environmental habitats.
Molecular methods based on direct bacterial nucleic acid extraction from environmental samples and subsequent amplification have the potential to overcome many restrictions of traditional microbiological approaches but are associated with other pitfalls (57). Efficient DNA isolation from soil is biased by incomplete cell lysis and nucleic acid adsorption to soil particles (13, 25, 31). Furthermore, soil-derived PCR inhibitors coextracted with nucleic acids affect downstream amplification reactions (38, 57). Although a multitude of DNA extraction and purification methods exist, including several commercially available kits, there is no universal standard protocol and methods have to be adapted to the specific experimental needs.
The present study was undertaken to develop a quantitative PCR method for the direct quantification of B. pseudomallei cells from soil. We first established a protocol for efficient DNA extraction and the removal of coextracted amplification inhibitors. B. pseudomallei cells were then detected as genome equivalents (GE) in a quantitative PCR (qPCR) using primers and a probe, specific to a 115-bp fragment of the type three secretion system 1 (TTSS1) of B. pseudomallei, which have recently been developed to detect this pathogen in both clinical samples and supernatants from environmental enrichment cultures (23, 24, 33). A nested PCR targeting a second B. pseudomallei-specific sequence (52) was applied to qualitatively confirm the qPCR results. This experimental approach was validated on 40 environmental soil samples collected in Northeast Thailand and proved to be a highly sensitive tool for rapid environmental surveillance of B. pseudomallei, compared to culture methods.
Bacterial strains used are listed in Table 1. Bacteria stored in frozen vials were streaked onto Columbia agar plates supplemented with 5% sheep blood (Becton Dickinson, Heidelberg, Germany). The plates were then incubated overnight at 37°C in air. Fresh cultures grown on agar were harvested for bacterial DNA extraction.
The PCR methods were validated on sandy loam soil samples collected in January (samples S 01 to S 95) or March (samples RFT 01 to RFT 30) 2010 from randomly selected rice fields with known high positivity rates for B. pseudomallei in Amphoe Lao Sua Kok, Ubon Ratchathani province, Northeast Thailand, as described previously (56). Briefly, a standard soil sampling technique was used (29), and 200 g of soil was collected from a depth of 30 cm. Soil samples sealed in plastic bags and plastic containers were shipped by air freight at ambient temperatures to the laboratory in Greifswald, Germany, and stored at room temperature until analyzed, because low temperatures are able to reduce the viable cell count of B. pseudomallei (55). Samples S 01 to S 95 and RFT 01 to RFT 30 were processed for DNA extraction (see below) within 10 days after soil sampling in Northeast Thailand. There was no significant difference in the qPCR-based B. pseudomallei detection (see below) between subsamples extracted on different days (data not shown). The original 200-g soil samples were mechanically homogenized prior to the experiments, and three 1-g, 0.5-g, or 0.1-g subsamples of each original 200-g sample were processed for DNA extraction and purification using different methods as described below. For culture experiments, three 25-g subsamples were processed as previously described (56). Additionally, 10 soil samples were taken from a 30-cm depth from agricultural land around Greifswald, Germany, as negative controls from an area of nonendemicity to confirm the specificity of the PCR methods and to test for cross-contamination.
The single PCR data of the 40 soil samples from this study were compared to the single culture results of the same samples, previously reported as summarized data by Trung and colleagues (56). Briefly, B. pseudomallei cells were detached from the soil matrices of 25-g subsamples by shaking in 50 ml of a solution containing 2.5% (wt/vol) polyethylene glycol 6000 (SERVA Electrophoresis, Heidelberg, Germany) and 0.1% (wt/vol) sodium deoxycholate (Merck KGaA, Darmstadt, Germany) for 2 h. The soil particles were then sedimented by centrifugation at 1,400 × g for 10 min. One hundred microliters of the supernatant was used to enumerate B. pseudomallei cells on Ashdown's agar (containing 10 g of Trypticase soy broth, 40 ml of glycerol, 5 ml of 0.1% crystal violet, 5 ml of 1% neutral red, 5 mg of gentamicin, and 15 g of agar per liter). For a qualitative culture, 1 ml of the same supernatant was mixed with 9 ml of Galimand broth (threonine-basal salt plus colistin [TBSS-C50]) (61), supplemented with 1 g/liter of polymyxin B. Enrichment cultures were incubated at 40°C for 4 days before being plated on Ashdown's agar plates. All plates were incubated at 40°C for 4 days, and the numbers of B. pseudomallei CFU were determined.
Total genomic DNA was extracted from mechanically homogenized soil samples by two commercially available kits according to the manufacturers' instructions, except where noted, and by the extraction method developed in this study.
The SoilMaster DNA extraction kit (Epicentre Biotechnologies, Madison, WI) was used according to the manufacturer's instructions. Briefly, 0.1 g of soil was added to 250 μl of soil DNA extraction buffer. Then bacterial cell lysis and DNA purification were performed using a hot detergent lysis process and a column chromatography step, respectively. The final precipitated DNA preparation was resuspended in 50 μl of Tris-EDTA (TE) buffer and stored at −20°C until used.
The FastDNA spin kit for soil (MP Biomedicals, Illkirch, France) was applied according to the manufacturer's instructions. Briefly, 0.5 g of soil was added to the lysing matrix A tube, which contained 987 μl of sodium phosphate buffer, 122 μl of MT buffer (from the Fast DNA spin kit), and a mixture of ceramic and silica particles. Then the bacterial cell disruption and DNA purification steps were performed using a vortex and a silica-based procedure, respectively. The final DNA preparation was eluted in 50 μl of DNase–pyrogen-free water and stored at −20°C until used.
The DNA extraction part of the protocol developed in this study is based on the protocol of Gabor et al. (14), with slight modifications. First, 1 g of soil was mixed with 750 μl of lysis buffer (100 mM Tris HCl, 100 mM Na2EDTA, 1.5 M NaCl, 1% cetyltrimethyl ammonium bromide [CTAB], pH 8.0) by vortexing the mixture at maximal speed until the soil was completely homogenized. After the addition of 40 μl of lysozyme (50 mg ml−1) and 10 μl of proteinase K (10 mg ml−1), the tubes were incubated at 37°C for 30 min. Then 200 μl of SDS (20% wt/vol) was added prior to incubation at 65°C for 2 h, with a vigorous vortex for several seconds after 1 h of incubation. The first supernatant (supernatant A) was collected by centrifugation at 6,000 × g for 10 min, and the soil pellet was reextracted by adding 1 ml of lysis buffer, in contrast to the original protocol (14). Vortexing for a few seconds was followed by incubation at 65°C for 30 min. Centrifugation was performed at 6,000 × g for 10 min, and the second supernatant (supernatant B) was collected and combined with supernatant A. Instead of chloroform (14), an equal volume of chloroform-isoamyl alcohol (49:1) was added to the mixture before the DNA was precipitated from the upper water phase by an addition of 0.6 volume of isopropanol and incubated at −20°C overnight (or at least 1 h). The DNA precipitate was collected by centrifugation at 16,000 × g for 7 min, washed with prechilled 70% ethanol, and resuspended in 50 μl of TE buffer, followed by incubation at room temperature for 1 h.
To purify DNA from potential coextracted PCR inhibitors, a modified form of Moreira's protocol (35) was used. Crude soil DNA was gently mixed with an equal amount of melted 1.6% low-melting-point (LMP) agarose (AppliChem, Darmstadt, Germany) in a sterile 2-ml tube. After the tube was allowed to stand at 4°C for 15 min, the solidified DNA-agarose matrix was subjected to three washing steps, with each step performed by adding 1.5 ml of TE buffer to the tube and placing the tube horizontally at 4°C for 24 h. After the last washing step, the purified DNA remained embedded in the LMP agarose matrix and was stored at −20°C. Before incorporation into the PCR, the DNA-agarose block was incubated at 70°C for 2 min and the melted solution was used as a template. Initial experiments revealed that the melted agarose solution did not affect PCR performance (data not shown).
A quantitative PCR assay was developed in order to detect the presence of PCR inhibitors coextracted with nucleic acids in the soil DNA extracts and to validate the quality of the different soil DNA extraction methods. The plasmid pCR2.1-IAC was constructed by inserting an artificial synthesized DNA fragment (MWG-Biotech, Ebersberg, Germany) of 135 bp (AGCCGACACGCGTCTCTATACTGTCGAGCAATCGGCGGATATCCCGTCACGCTGTTTGTGATCGGCGTTATCGCGTTCTTGA TCGCACTTTACGAAGCTGTTACGGATACTGACCGGTGTGCACGCGG GCGCGCA), containing a unique nucleotide sequence of cytomegalovirus (indicated in bold), into a pCR2.1 plasmid (Invitrogen, Darmstadt, Germany). The quantitative PCR assay of the recombinant plasmid pCR2.1-IAC had the same thermal conditions as the TTSS1 target amplification. One hundred copies of pCR2.1-IAC were incorporated into a 25-μl reaction mixture as described for the following TTSS1 gene qPCR assay (see below), except that the primers and the probe were replaced with a 400 nM concentration each of primers IAC2 forward and IAC2 reverse and a 260 nM concentration of CMV3 probe (MWG-Biotech, Ebersberg, Germany), labeled with hexachloro-6-carboxyfluorescein (HEX) at its 5′ end and black hole quencher 2 (BHQ2) at its 3′ position (Table 2). The thermal cycling profile was as described for the TTSS1 qPCR assay (see below), except that the number of thermal cycles was set to 60. The degree of PCR inhibition caused by soil DNA extracts was determined by cycle threshold differences (ΔCT), which were calculated by subtracting the CT values of the PCR with the recombinant plasmid and soil DNA extracts from the CT values of the PCR mixture containing the recombinant plasmid as a single template (control).
The TTSS1 gene qPCR mixture, at a final volume of 25 μl, consisted of 1× TaqMan universal PCR master mix (Applied Biosystem, Branchburg, NJ), which contained the following: AmpliTaq Gold DNA polymerase and AmpErase uracil-N-glycosylase (UNG); a 400 nM concentration each of primer BpTT4176 forward and BpTT4290 reverse; a 260 nM concentration of BpTT4208 probe (MWG-Biotech, Ebersberg, Germany), labeled with 6-carboxyfluorescein (FAM) at its 5′ end and black hole quencher 1 (BHQ1) at its 3′ position (Table 2); 10 μg of nonacetylated bovine serum albumin (BSA; Sigma-Aldrich, St. Louis, MO); and 4 μl of purified soil DNA embedded in an agarose matrix as a template. Amplification and detection were performed on the Mx3000P qPCR system (Stratagene, Cedar Creek, TX) using the manufacturer's standard settings. Thermal conditions were 50°C for 2 min to activate UNG, which prevents carryover contamination, followed by an initial denaturation step at 95°C for 10 min and 45 cycles of denaturation at 95°C for 15 s and amplification at 60°C for 1 min. The cycle threshold (CT) values were automatically calculated by applying adaptive baseline algorithms (MXPro-Mx3000P v. 320, build 340). The primers used were empirically tested and produced no artifacts (data not shown).
To further improve the sensitivity of B. pseudomallei detection and to reduce the effect of any remaining PCR inhibitors, a nested-PCR approach was applied. The outer primers, 174 forward (174F) and 725 reverse (725R) (Table 2), were used to amplify a 552-bp fragment of BPSS1187, which encodes a hypothetical B. pseudomallei-specific protein (according to the genome sequence of B. pseudomallei K96243) (52). The first PCR was performed in a 25-μl reaction mixture which consisted of 1× PCR buffer containing 1.5 mM MgCl2, 1 U of AmpliTaq Gold DNA polymerase (Applied Biosystem, Foster City, CA), a 125 μM concentration each of deoxynucleotides dATP, dCTP, dGTP, and dTTP (Roche, Mannheim, Germany), a 400 nM concentration each of primers 174F and 725R (Table 2), 10 μg of BSA, and 4 μl of purified soil DNA immobilized in solid agarose as a template. Thermal cycling was carried out in an Uno II thermocycler (Biometra, Goettingen, Germany), with an initial denaturation step of 95°C for 5 min, followed by 35 cycles of 95°C for 30 s, 60°C for 30 s, and 72°C for 45 s, and a final extension step at 72°C for 5 min. One microliter of the resulting PCR product was then applied as the template in a second PCR using the 8563 PCR assay (Table 2) as described by Supaprom and colleagues (52). A melting-curve analysis was performed using the SYBR green I master kit (Roche Applied Science, Mannheim, Germany) to check for primer-dimer artifact formation.
To determine the detection limits and efficiencies of the TTSS1 qPCR genomic DNA from freshly grown cultures of B. pseudomallei, B. pseudomallei strain K96243 was isolated using the DNeasy blood and tissue kit (Qiagen, Hilden, Germany). Quantification was performed using SYBR green I master mix (Stratagene, Cedar Creek, TX) and lambda DNA as the standard (molecular weight marker XV; Roche, Mannheim, Germany), by following the manufacturer's protocol, on the Mx3000P qPCR system (Stratagene, Cedar Creek, TX). The amount of DNA was then converted to genome equivalent (GE) copies based on the B. pseudomallei K96243 genome size of 7.25 Mb (17):
For the detection of B. pseudomallei in soil samples by using either TTSS1 qPCR or BPSS1187 nested PCR, each of the three 1-g subsamples was analyzed in duplicate. TTSS1 qPCR bacterial counts of each replicate, expressed as genome equivalents (GE) per PCR mixture, were determined with the standard curve shown in Fig. 1. The bacterial count per g of soil was calculated with the following equation:
The statistical significance of differences was determined by two-tailed Student's t tests and Wilcoxon signed rank tests with GraphPad Prism software version 4.0 for Windows (GraphPad Software, San Diego, CA). The quartile coefficient of dispersion (interquartile range/median) was used to specify the intrasample variation of qPCR results obtained in the three PCR runs, each performed in duplicate.
We first evaluated the quality of genomic DNA with respect to coisolated PCR inhibitors after using different extraction methods in 10 soil samples from Northeast Thailand. These samples were culture positive for B. pseudomallei. The genomic DNA was isolated with either the SoilMaster DNA extraction kit, the FastDNA spin kit for soil, or our method based on enzymatic and chemical methods plus subsequent purification in an agarose matrix. A noncompetitive internal amplification control was applied to determine the rate of inhibition. The differences in cycle threshold (ΔCT) values between PCRs with and without soil DNA extracts, supplemented with 100 recombinant plasmid copies, were used to determine the grade of inhibition. All three tested DNA isolation methods failed to completely remove PCR inhibitors coisolated with genomic DNA from soil. Table 3 (columns without BSA) shows that accurate detection of the copy numbers of the incorporated recombinant plasmid failed for 6 samples (60%) isolated by the SoilMaster DNA extraction kit, with either no amplification at all or very high ΔCT values, and for all 10 samples (100%) obtained from both the FastDNA spin kit for soil and our developed protocol. The majority of soil DNA extracts showing no qPCR amplifications were brownish in color, indicating the presence of coextracted soil compounds interfering with the qPCR assay. Additional washing steps using either 5.5 M guanidine thiocyanate or humic acid wash solution, optional in the FastDNA spin kit, did not improve PCR performance for DNA samples isolated by this kit (data not shown).
We overcame PCR inhibition by the incorporation of nonacetylated BSA into the qPCR assay as follows. All 26 samples that were initially qPCR negative displayed amplification in the presence of BSA (Table 3, columns with BSA). The ΔCT values for each of the three tested extraction protocols, with 10 soil samples each, were significantly reduced using BSA as a PCR adjuvant, with overall ΔCT values of 0.48 ± 0.09, 0.47 ± 0.22, and 0.85 ± 0.17 for the SoilMaster DNA extraction kit, the FastDNA spin kit for soil, and our developed protocol, respectively (Table 3, columns with BSA).
We then optimized a TTSS1 gene qPCR assay which should subsequently be used for the direct quantitative detection of B. pseudomallei in soil. Figure 1 shows the straight calibration line relating the CT values to the numbers of B. pseudomallei genome equivalents by using a 10-fold dilution series of genomic B. pseudomallei K96243 DNA. There was a strong linear inverse relationship (r2 = 99.9%) between the CT values and the log10 genome equivalents of B. pseudomallei over 6 orders of magnitude. Higher variations of the CT values were observed if fewer than 10 genome equivalents per PCR mixture were used. Therefore, 10 B. pseudomallei genome equivalents per PCR mixture was used as the lower limit for calculation. The limit of detection (LOD) was 3 B. pseudomallei genome equivalents per single qPCR mixture, corresponding to 75 B. pseudomallei cells per g of soil when assuming a DNA extraction efficiency of 100%.
The efficiency of qPCR amplification was 98.7%. To ensure accurate quantification of the qPCR, melting-curve analyses were conducted, which confirmed the absence of any primer-dimers (data not shown).
To qualitatively confirm any TTSS1-based qPCR signal by a second PCR target in case of negative results in culturing, we also established a BPSS1187 gene-based nested PCR. The LOD of this assay was 1 GE/PCR mixture with a used volume of 5 μl. This corresponds to a theoretical value of 20 GE per g of soil for the qualitative detection of B. pseudomallei. The specificity of the TTSS1 primers and probe and the specificity of the inner BPSS1187 primers and probe for B. pseudomallei strains have already been rigorously tested in previous studies (24, 37, 52). In accordance with these results, our extended testing of the primers and probes of both PCR methods against 73 isolates of species that are closely related phylogenetically, including species such as Burkholderia seminalis, Burkholderia latens, Burkholderia diffusa, and Burkholderia pyrrocinia (listed in Table 1; data not shown), revealed no PCR amplification, whereas all 29 different B. pseudomallei isolates proved to be positive in our assays.
To finally select the DNA extraction protocol to be used for the qPCR-based quantification of B. pseudomallei in soil, we compared the results of the TTSS1 qPCR with added BSA for five DNA soil samples (S 03 to S 07) processed by the three different DNA extraction methods. Despite the lower average ΔCT values obtained for the two commercially available kits, our newly developed method led to a higher detection rate of B. pseudomallei genome equivalents, indicating a better template quality or quantity (Fig. 2). The detection factors for the SoilMaster DNA extraction kit and FastDNA spin kit for soil were 0.24 ± 0.01 Times and 0.52 ± 0.05 times that of our method, respectively.
Our PCR systems were validated using 40 environmental soil samples collected in Northeast Thailand. The same samples were used in a parallel study in which we validated our newly developed protocol for the culture-based detection of B. pseudomallei based on soil dispersion in a polyethylene glycol and sodium deoxycholate solution (56). In addition, we tested 10 soil samples from Germany as negative controls from an area of nonendemicity. A sample was classified as PCR positive when B. pseudomallei DNA could be detected in at least one replicate of the three 1-g subsamples. The results in Fig. 3 show that all 26 samples from Northeast Thailand which were B. pseudomallei positive by direct quantitative culture were B. pseudomallei qPCR positive (Fig. 3A, 13 samples with >25,000 GE/g of soil; Fig. 3B, 13 samples with 300 to 25,000 GE/g of soil), with a median of 1.84 × 104 genome equivalents per gram of soil (range, 3.65 × 102 to 7.85 × 105). In those 26 samples, raw data of colony counts were poorly linearly related to qPCR results (r2 = 0.629), whereas log transformation of culture and qPCR data resulted in a high correlation (r2 = 0.96). All samples exhibited a ΔCT value below 2 (data not shown), indicating no significant inhibition of qPCR amplification. Moreover, determination of total DNA by fluorescent dye staining or of total bacterial DNA by bacterial 16S rRNA gene copies from representative soil samples did not correlate with the quantity of B. pseudomallei GE (data not shown), indicating that the degree of B. pseudomallei DNA detection was not generally linked to the amount of bacterial DNA present or to the efficiency of recovering bacterial DNA from single samples.
Although described almost a century ago (60), the worldwide environmental distribution of B. pseudomallei is still unknown and our understanding of the environmental factors determining the presence of B. pseudomallei is rudimentary. As a basis for a better understanding of B. pseudomallei ecology in its natural habitat, quantitative culture-dependent and quantitative culture-independent molecular methods are needed to detect the organism. The two methodological approaches are likely to complement rather than replace each other. To the best of our knowledge, the direct quantitative detection of B. pseudomallei from an environmental habitat using molecular methods has not been reported. We therefore aimed to establish a quantitative DNA-based method to detect B. pseudomallei in soil samples from an area of endemicity.
The soil samples used for validation of our PCR methods originated from randomly selected rice fields in Amphoe Lao Sua Kok, Ubon Ratchathani province, Northeast Thailand, and consisted of sandy loam taken at a depth of 30 cm. This habitat is normally anoxic except for the rhizosphere of the rice plants (27), where members of the order Burkholderiales can also be found (30). Generally, Proteobacteria represent a minor group of the whole microbial community in bulk rice field paddies (27).
Direct molecular bacterial detection from soil is methodologically challenging because PCR inhibitors are often coextracted with the DNA. Depending on the soil type, different concentrations of inhibitory components can be found. These include humic acid, fulvic acid, polysaccharides, and metal ions (59), all of which negatively affect the DNA polymerase activity and/or the availability of DNA templates (38). Various strategies have been proposed for excluding or reducing inhibitory effects in soil samples prior to PCR, such as the use of cesium chloride density gradient ultracentrifugation (4), dialysis (3), cetyltrimethylammonium bromide (CTAB) for complexing inhibitors, polyvinylpolypyrrolidone (PVPP) (63), chromatography, electrophoresis, or multivalent cations (59) or the procedures of separating inhibitors from nucleic acids applying gel filtration (34), washing DNA immobilized in agarose (35), or using PCR additives, such as nonacetylated bovine serum albumin (BSA) and phage T4 gene 32 protein (59).
PCR inhibition was also observed in this study for the two commercial soil DNA isolation kits and the protocol developed in this study. Although both commercial kits tested coupled DNA extraction with purification by either chromatography (SoilMaster DNA extraction kit) or silica (FastDNA spin kit for soil), inhibition of amplification was not significantly prevented until BSA was added to the PCR reagent mix. This was also true for our protocol, which combined a conventional, slightly modified DNA extraction protocol with the washing of extracted DNA embedded in an agarose matrix. However, using this protocol together with BSA as a PCR additive resulted in the highest sensitivity for B. pseudomallei detection in soil with a TTSS1-based qPCR.
A limitation of our study is the restriction to soil samples of sandy loam only, which is the most prevalent soil type in Northeast Thailand. Therefore, further field studies have to demonstrate the general usefulness of our protocol for different soil types. For the calculation of the sensitivity of our PCR methods, we assumed a theoretical DNA extraction efficiency of 100%, being well aware that the soil type might influence the efficiency of bacterial DNA extraction. We addressed this issue in preliminary experiments in which we quantified total bacterial DNA by 16S rRNA gene copies in soil samples collected around Greifswald in Northeast Germany and in representative samples from Northeast Thailand. Our results revealed a comparable, even slightly higher bacterial load in the soil around Greifswald, including samples with clay loam- and silty loam-like textures (data not shown), indicating a potential usefulness of our protocol for other soil types.
Previous studies targeted the TTSS1 gene and BPSS1187 for the qualitative detection of B. pseudomallei in enrichment cultures of soil samples (24) and clinical samples (52), respectively. The results of our study confirm the specificity of these targets, since the TTSS1 gene and the BPSS1187 coding sequence were present in all B. pseudomallei bacteria tested. Furthermore, these sequences could not be detected within genomic DNA of closely related species. Importantly, the TTSS1 qPCR protocol developed in this study led to an improved PCR efficiency and a wider linear range of B. pseudomallei detection compared to those of a PCR protocol applied for the detection of B. pseudomallei in environmental enrichment cultures, with the same TTSS1 gene sequence used as the target (24).
Combining the TTSS1 qPCR assay with the BPSS1187 gene nested approach, we could classify 35 out of 40 soil samples as B. pseudomallei positive. Out of these B. pseudomallei-positive samples, 26 samples were positive using our recently improved quantitative culture-based method (56) and therefore were directly compared to our qPCR protocol developed in this study. The significantly higher numbers of B. pseudomallei bacteria detected in those 26 soil samples by TTSS1 qPCR (Fig. 3) might be explained by the coextraction of extracellular DNA and/or DNA originating from viable but nonculturable cells of the bacterium (18). Although the subsamples used for PCR consisted of much less soil material than the subsamples used for the culture method, the TTSS1 qPCR resulted in a significantly lower intrasample dispersion. It seems likely that an uneven distribution in the culturable proportion of B. pseudomallei populations, which might be due to differences in the hydration status of the soil as well as in the soil type itself (54), is responsible for this phenomenon.
Thirty (96.8%) out of 31 samples which were positive by either direct culture or enrichment culture resulted in a positive signal in the TTSS1 qPCR assay. The only exception was sample RFT 23, in which case the culture after the enrichment step was positive for B. pseudomallei but the TTSS1 gene amplification was negative. However, this sample tested positive in the nested BPSS1187 PCR. Out of the 35 B. pseudomallei-positive samples, the TTSS1 qPCR detected more positive samples than did quantitative direct culture (33 [94%] versus 26 [74%], P = 0.045, Fisher's exact test). However, with our sample size, the qualitative sensitivities were not significantly different when we compared the 35 samples which were positive by either qPCR or nested BPSS1187 PCR to the 31 samples which were positive by either direct or enrichment culture (P = 0.39, Fisher's exact test).
In conclusion, our presented qPCR method is able to detect significantly larger numbers of B. pseudomallei bacteria within soil samples, with a lower dispersion of subsample results, than direct culturing methods. The nested-PCR approach detects B. pseudomallei in samples at a detection limit that is below that of the qPCR and thereby is able to further improve the culture-independent sensitivity of B. pseudomallei detection in soil. Taken together, our experimental system will likely help unravel the ecology of B. pseudomallei in its natural habitat.
We thank Gumphol Wongsuvan, Sukanya Pangmee, Premjit Amornchai, Sayan Langla, and Anne Krause for their excellent technical assistance.
This study was funded in part by the Wellcome Trust of Great Britain.
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Published ahead of print on 29 July 2011.
†The authors have paid a fee to allow immediate free access to this article.
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