Journal of Nucleic Acids

Journal of Nucleic Acids / 2013 / Article

Research Article | Open Access

Volume 2013 |Article ID 801505 | https://doi.org/10.1155/2013/801505

John P. Jakupciak, Jeffrey M. Wells, Richard J. Karalus, David R. Pawlowski, Jeffrey S. Lin, Andrew B. Feldman, "Population-Sequencing as a Biomarker of Burkholderia mallei and Burkholderia pseudomallei Evolution through Microbial Forensic Analysis", Journal of Nucleic Acids, vol. 2013, Article ID 801505, 13 pages, 2013. https://doi.org/10.1155/2013/801505

Population-Sequencing as a Biomarker of Burkholderia mallei and Burkholderia pseudomallei Evolution through Microbial Forensic Analysis

Academic Editor: Satoshi Obika
Received01 Aug 2013
Revised01 Oct 2013
Accepted02 Oct 2013
Published17 Dec 2013

Abstract

Large-scale genomics projects are identifying biomarkers to detect human disease. B. pseudomallei and B. mallei are two closely related select agents that cause melioidosis and glanders. Accurate characterization of metagenomic samples is dependent on accurate measurements of genetic variation between isolates with resolution down to strain level. Often single biomarker sensitivity is augmented by use of multiple or panels of biomarkers. In parallel with single biomarker validation, advances in DNA sequencing enable analysis of entire genomes in a single run: population-sequencing. Potentially, direct sequencing could be used to analyze an entire genome to serve as the biomarker for genome identification. However, genome variation and population diversity complicate use of direct sequencing, as well as differences caused by sample preparation protocols including sequencing artifacts and mistakes. As part of a Department of Homeland Security program in bacterial forensics, we examined how to implement whole genome sequencing (WGS) analysis as a judicially defensible forensic method for attributing microbial sample relatedness; and also to determine the strengths and limitations of whole genome sequence analysis in a forensics context. Herein, we demonstrate use of sequencing to provide genetic characterization of populations: direct sequencing of populations.

1. Introduction

Genome sequencing data of mixtures can function as biomarkers for identification of genetic content of samples and to establish a sample’s genome profile, inclusive of major and minor genome components, drill down to identify SNPs and mutation events, compare relatedness of genetic content between samples, profile-to-profile, and provide a probabilistic or statistical scoring confidence for sample attribution. While high-throughput, automated sequencing has been used for years, analysis of sequencing information has focused on consensus sequencing [15]. In addition, sequencing has been used to infer microbial relationships [68]. Due to the ease of generating large volumes of sequence data, there has been pressure to develop computational tools [9]. Novel approaches, based on probabilistic analysis of sequencing information for mixtures and metagenomic samples, enable a broad capture of sequence data from a single run to characterize multiple genomes in a sample, even in isolates that are considered pure [10, 11]. When identifying genomes and determining the distribution of related organisms, knowing the populations of genomes in a sample is critical to accurate biomarker detection of disease, especially to the strain-level of identification.

In this study, we investigated the capability of whole genome sequence analysis to characterize relatedness/association between closely related populations of known ancestry under controlled conditions in the laboratory. Accompanying goals were to verify whether whole genome sequencing analysis is a reliable microbial forensic method for attributing relatedness, characterizing the extent of evolutionary mutations of Burkholderia mallei (glanders) and Burkholderia pseudomallei (melioidosis) populations over time, and understanding the strengths and limitations of whole genome sequence analysis in a forensics context. Burkholderia pseudomallei is the causative agent of melioidosis. The 7.2 Mb genome of Burkholderia pseudomallei consists of two chromosomes of 4,074,542 bp and 3,173,005 bp [12, 13].

Overall, the knowledge of this pathogen and its disease is limited. Pulsed field gel electrophoresis (PFGE) and MLST have been the current standard for identifying strains of Burkholderia; however, multilocus variable number tandem repeat (VNTR) and MLVA methods have been introduced [1416]. The calculated mutation rate is mutations/generation, which is slightly greater than Y. pestis and E.coliO157 [17].

Burkholderia mallei is rarely associated with human infection, and is more commonly seen in domesticated animals such as horses, donkeys, and mules where it causes glanders. The B. mallei genome has undergone a substantial genome reduction since the last common ancestor with B. pseudomallei, facilitated apparently by recombination events at the highly identical copies of transposase genes that are widespread in the B. mallei genome. The pathogen is hostadapted and is not found typically in the environment outside of its host [18]. Glanders is often fatal if not treated with antibiotics, and transmission can occur through the air, or more commonly when in contact with infected animals. Rapid-onset pneumonia, bacteremia (spread of the organism through the blood), pustules, and death are common outcomes during infection. The virulence mechanisms are not well understood [19, 20].

The Gram-negative pathogen B. pseudomallei is often fatal disease of both humans and animals [21]. Endemic to parts of South East Asia and Northern Australia, it is considered a major tropical pathogen and Category B biowarfare agent [22, 23]. In many countries, B. pseudomallei can only be experimentally manipulated under biosafety level 3 (BSL3) conditions.

1.1. Background of Microbial Forensics

Determining the source population of a biothreat agent attack is a goal of microbial forensics. Forensic scientists attempt to uncover DNA evidence to pinpoint a suspect and verify evidence for proof of guilt that will hold up in court. Microbial forensics played a major role in the Amerithrax investigation following the 2001 anthrax attacks on members of Congress and the public [24]. The utility of direct sequencing analysis for characterization of sample populations is a new paradigm for genome identification [11]. Recent investigations have shown strategies to process high-throughput DNA sequencing for accurate identification of major and minor populations and their concentrations [10].

For many years, microbial forensics relied heavily on amplified fragment length polymorphism (AFLP) and polymerase chain reaction (PCR) methods to attribute “relatedness” between crime scene evidence and suspected source samples [25]. After validation of hundreds of thousands of probes, high density microarrays can detect expected genomes, based on probe selection [26]. Since the emergence of next generation sequencing platforms within the last few years, genomic analysis of whole genome data is gaining popularity as a forensic technique. Whole genome sequencing provides vastly more quantity and specificity than either PCR or AFLP, and the implementation, reliability, and limitations of microbial genomic analysis need to be optimized, validated, and understood.

A number of studies combined AFLP, restriction fragment length polymorphism (RFLP), and PCR techniques for better discrimination between bacterial pathogen strains. AFLP involves cleaving sample genomic DNA with two or more restriction enzymes at base specific positions, then analyzing changes in DNA fragment lengths between different samples [2729]. Changes in fragment length indicate changes in base identities or indels at the points where the restriction enzymes cleave the genomic DNA. Each fragment length change becomes a biomarker that may be compared with other strain biomarkers to construct a phylogenetic relationship among the separate populations.

Early PCR analysis of pathogenic bacteria was achieved by as few as 2–4 primers 20–24 nt in length [30]. As PCR techniques advanced, multiplexing primers were used to analyze various known static biomarkers that discriminate between different microbial pathogens [31]. Modern PCR arrays can achieve over 100 fold oligomer target specificities [32]. Even 100-fold multiplexing of oligomer targets represents a small proportion of the available number of biomarkers that can be detected with WGS. Furthermore, only short, predetermined primer sequences are detected with PCR, requiring prior knowledge about the sample for satisfactory primer selection. In addition some targets are shared across species. For example, screening of O-antigen genes showed the presence of multiple O-antigen types and sero-cross reactivity in near-neighbor species [33].

On the other hand, PCR amplifies minute amounts of DNA within a much larger background signal. Thus, rare variant polymorphisms or low concentration pathogens may be detected with PCR, if the right primers are available. Due to its amplification ability in the presence of high background, long PCR amplicons will likely continue to have a purpose in microbial forensic analysis for the detection and validation of rare variant genomes mixed in populations [34]. The limitations of conventional methods have been reviewed elsewhere. The focus of this paper is on the use of sequencing to characterize genome populations in a sample. Population-sequencing is an economic way of assessing species diversity that provides resolution at the strain level [3540].

The first bacterial genome sequences were published in 1995, and within 15 years, over a thousand fully sequenced bacterial genomes have become publicly available [41].

Using sequencing to measure relative populations to identify simultaneously the organisms present in microbial populations with respect to the specific taxa (e.g., genus, species, subspecies, and strain) of bacteria, viruses, parasites, fungi, or nucleic acid fragments including plasmids and mobile genomic components requires bioinformatics tools to accurately match sequencing output. The greatest limitation of WGS is the inherent machine errors arising from the DNA sequencing process. Commonly reported sequencing error rates range from 0.1% for Illumina sequencers to 15% (single pass error) for PacBio single molecule sequencers [42]. Even at the level of a 0.1% error rate, the actual presence of a sequence variation in sample data must be verified statistically. For example, for a genome maintained in GenBank that has approximately 3 million base pairs, even, assuming, the best case scenario sequencing error rate of 0.1%, will generate 3,000+ errors spread throughout the entire genome [43].

Erroneous sequences appear identical to true variations within the data. Therefore, variant validation, or a calibrant [11] is required before trusting sequence variations reported by the sequencer. Stringency conditions applied to variant validation include(i)variants must appear in both forward and reverse sequenced reads,(ii)minimum percentage of consensus,(iii)minimum number of occurrences in agreement,(iv)base calling quality score.

To distinguish between errors and true positive calls, Bayesian statistics are applied to calculate posterior probabilities for the validity of each variant detected. Optimally, each criterion can be used to enhance or diminish the probability for each variant’s trustworthiness. In addition, given the rate of genomic mutation and growing evidence of horizontal gene transfer, static methods that rely on predetermined signatures produce false negative results if (a) mutation has occurred in the nucleic acid sequence of the sample relative to signatures, (b) the target signature was horizontally transferred, or (c) genomic near neighbors are present in the sample.

Our approach involved a combination of alignment and alignment-free comparisons of sequence information. Our genomic sequence data involved massive amounts of relatively short reads (~100 nt). For genome assembly, it would be more ideal to sequence sample DNA with a platform that provides long read or paired end data for better assembly of large (supercontigs). Several groups, including our own, also utilize unaligned, unassembled sequence approaches to analyze genomic data [44]. Advantages of unaligned approaches include(i)the ability to examine all reads rather than the subset of aligned reads,(ii)faster results that require less expert analysis,(iii)more straightforward scoring for genome matches.

On the other hand, aligning reads against reference genomes enables the extraction of genomic details not available from unaligned approaches. Such details include(i)longer length matches to reference sequences,(ii)varying gene recombinations and indels within the genome,(iii)better discrimination between similar sequences,(iv)determination of coverage depth and percent genome coverage,(v)more accurate detection and validation of polymorphisms.

2. Materials and Methods

2.1. Experimental Design

The purpose of this experiment was threefold as follows.(1)Observe DNA sequence mutations arising from an originally clonal isolate of Burkholderia mallei strain China 7 (ATCC 23744), and Burkholderia pseudomallei strain 668 when cultured under stressful conditions in the laboratory.(2)Determine the strengths and limitations of whole genome sequence analysis for characterizing variation between similar substrains.(3)Advance methods for determining “relatedness” between microbial samples.

In the forensics use case, evolutionary pressure and manipulation in the laboratory can be measured to assess dynamics of bacterial metapopulations. We incorporated stress in our experiments because we wanted enough variation under laboratory conditions to compare lineages to source to assess WGS utility in attribution. For example, samples maintained under controlled conditions, laboratory stocks, may show less diversity than samples found in nature. Instead of analysis at a few stable sites, we evaluated analysis across the entire genome and to use the entire genome as a biomarker to build genome population content (context) profiles of samples to maximize data for microbial forensics to establish statements regarding the relationship of manipulated samples and reference samples.

The sequences were from a single colony, which corresponds to a single constituent within the parent population. Generally, during growth, this bacterium will accrue new random, and fitness-selected mutations, as well as rearrangements that result in consensus and population differences in the sequence content with respect to the reference genome.

A single colony of Burkholderia pseudomallei and Burkholderia mallei was passaged into 12 different plates. These twelve bacterial cultures were maintained separately over the course of seven more passages. Each culture passage was started with a single clonal colony. This created a single genome bottleneck at each passage step. Mutational variations differentiating each lineage were thus a result of initial variation in the source clonally derived culture plus mutations accumulated during the course of the eight growth and passage steps (Figure 1).

There are two approaches that may be taken relative to the genetic bottlenecks at each passage step. The first approach, which we chose for this experiment, involved reducing all intralineage variation down to approximately zero by selecting only one clonal colony for passaging. The second approach would be to randomly select a subset of cells from the flask for subsequent passage. This approach should theoretically maintain a higher level of genetic diversity among the “lineage,” since not all microbes would be related to each other within each lineage at the end of the experiment. As genetic mutations accumulate over many generations, each lineage theoretically should become more diverse within itself; however, many of the cells within the same lineage would be no more related to each other than they would be to microbes in another lineage (Figure 2).

At the end of eight passages, six clones from each of the twelve lineages were collected. DNA was isolated from each clone and sequenced using an Illumina Genome Analyzer IIx platform as follows:(i)single ended,(ii)average read length,(iii)other read statistics.

2.2. Culturing Media and Methods

Burkholderia mallei and B. pseudomallei were alternately cultured in tryptic soy broth, followed by culturing on selective PC agar plates for a total of seven (7) passages. Bacterial DNA was isolated from the eighth passage liquid culture. At each subsequent solid media passage step, the liquid culture was plated to yield single colonies. A single colony was selected to inoculate the next liquid media passage. A portion of the colony selected for passage, along with five (5) additional colonies from each of the twelve cultures, was frozen and stored as an archive for potential future analysis (Figure 3).

2.3. DNA Sequencing Protocol

Single ended short read DNA sequencing was performed using a Genome Analyzer IIx (GA IIx) (Illumina, San Diego, CA, USA). Library preparation was performed using a genomic DNA sample preparation Kit. DNA clusters were generated according to the manufacturer’s instructions using an Illumina cluster generation kit (Multiplexing Sequencing Primers and PhiX Control Kit v2) on an Illumina cluster station. All sequencing runs were performed with the GA IIx using the Illumina TruSeq SBS kit v5. Fluorescent images were analyzed with the Illumina CASAVA 1.8 software to obtain FASTQ-formatted sequence data of the short reads. For further details of the experimental methods, see Eppinger et al. [45].

2.4. Identification and Characterization of Major and Minor Components

We evaluated algorithms to process millions of short sequence reads from complex community metagenomic data. Orthogonal read alignment tools were used for comparison, GNUMap/SOAP (pipeline A), and pipeline B.

2.5. Selection of Assembly Software

SOAP implements the algorithm (Burrows-Wheeler transform), whereas GNUmap uses a probabilistic Needleman-Wunsch algorithm, which takes advantage of Illumina probability files to improve the mapping accuracy for lower quality reads and increase the amount of usable data produced in a given experiment. SNP calling was done under a Bayesian Inference framework as implemented in SOAPsnp by comparing an assembled genome against the reference genome used. The output files of the pipeline are a nucleotide fasta file per chromosome, a GenBank annotated file per chromosome, and a SNP file with information on all chromosomes. Performed alignment and variant analysis (SNPs and other mutations). Identified SNPs and compared relatedness of clones by building phylogenetic trees using trusted SNP variations. Trusted SNPs must have posterior probabilities >50% (Table 1).


123456789101112131415

AT36*T3744A36337110
AG99G3866A0006110

This example SNP is called with a posterior probability of 36%. The reference genotype is A and the consensus is T. The best base identified is T with a -score of 37, while the 2nd best base is the same as reference with a -score of 36. The total depth at the site is seven, with four reads supporting T and three reads supporting A. This SNP should be rejected by setting an appropriate posterior probability cutoff and not used to determine diversity or phylogenetic distance relationships.
2.6. SNP Accuracy and Depth of Coverage

We compared the depth of coverage in terms of the number of reads covering a particular SNP and compared that depth of coverage to the resulting posterior probability () of the SNP read. The resulting analysis shows that there is a general trend of increasing with increasing depth.

2.7. Phylogenetic and Diversity Estimates

Phylogenetic relationships were estimated via both Maximum Likelihood (RAxML) and a network approach (SplitsTree) to estimate relationships among strains. Distance measurements are based on SNPs with posterior probabilities <95% for chromosome one and chromosome two (Figure 4).

2.8. Phylogenetics Is Assessed by Measurements of SNPs through Lineages

SOAPsnp takes into account base calling quality scores, as well as the number and percentage of reads receiving a particular base call at that position when calculating posterior probability scores. Theoretically, the lowest posterior probability score possible for the called base is 25% for equal probability among all bases. Posterior probabilities below 50% for an individual clone’s SNP should not be trusted, unless supported by presence of the same base SNP in another clone of the same lineage.

The percent consensus column indicates consensus between accepted base calls among the different clones at that potential SNP position. The average posterior probability column indicates the average probability among clones with the particular called SNP. The Maximum Probability is an upper bound of probability that a particular SNP occurs in at least one of the five clones. It is calculated by where is the first posterior probability for a called SNP, is the second posterior probability, and is the number of clones possessing the called SNP. is the probability of a condition not being met for data set “,” thus if there is an 80% chance of meeting an AND/OR condition in data set 1, the remaining 20% chance is multiplied by the chance of not meeting a condition in data set 2, multiplied by the percent chance of not meeting the condition in the next set and so on.

We set a SNP trusting threshold at 95% probability. If the probability values are ≥80% but <95, then it is considered probable that the SNP is present and not used in constructing phylogenies for attributing relatedness among samples or diversity.

Pipeline B. We trained this pipeline on in-silico data to validate algorithm performance on being able to distinguish bacteria down to strain level and to differentiate between highly related genomes. The process was performed in blinded fashion and genome identification was based on probabilistic values for matching criteria. Probability of obtaining matches above threshold of established criteria needs to be calculated for each matched position. The level of confidence is based on variations from reference genome sequences. Data sets possessing similar sequence variations are clustered together by phylogenetic analysis.

Pipeline B is an integrated set of public domain tools and a custom SNP calling method modified from the BFAST algorithm to find a candidate alignment position for each read. The mapping depends on a set of index masks to determine which locations in a read require matching as part of the scoring process. The aligned reads are converted to the public domain SAM format, sorted according to position along the reference and then submitted for mPileUP analysis via the Samtools software suite [11].

3. Results and Discussion

Sequence reads were trimmed and filtered by a custom script. Orthogonal read alignment tools were used for comparison, GNUMap/SOAP (A) and pipeline B. Figure 5 displays our genomic data analysis pipeline A. GNUMap is better at detecting reads that differ from the reference sequence. SOAP is much faster at aligning raw reads than GNUMap, but GNUMap attains a higher percentage of aligned reads than SOAP. The differences in performance can partially be explained as speed versus accuracy tradeoffs between heuristic and probabilistic algorithms. Most other approaches such as search engines like BLAST rely on heuristic analysis to find approximate answers to questions. Heuristic approaches, however, are limited because they only identify simple alignments or lack of alignments in data, and do not guarantee that all possible matches are detected. Furthermore, since we observe a continuum of variants, the concept of characterization of samples based solely by SNP calls becomes somewhat perilous from a forensic point of view. The population structure within a single colony may arise from multiple factors, such as growth time and conditions. While we are not aware of specific quantitative analyses of how total culture time impacts population diversity, it is conceivable that the number of SNPs called based on the fraction exceeding 0.5 in frequency could indeed depend on such factors. This makes standardization of protocols a high priority for comparative analysis requiring growth in culture. It would be most advantageous to avoid any culturing step when there is sufficient available sample for direct analysis by NGS, but this, of course, cannot be guaranteed in practice.

In Table 2, we summarize specific Pipeline B alignment results for Burkholderia pseudomallei and Burkholderia mallei and Bacillus globigii for comparison to our nonalignment screening analysis results. As discussed above, two Burkholderia species had statistically high values outside the expected range for an exact match. These nonalignment results can be explained by data in the third column in the table, which shows the total fraction of nonreference base calls for reads successfully aligned to the reference genome. For Bacillus globigii, the baseline value of 0.018 reflects the contributions of the base calling and indel error rate of sequencing process, as well as population structure within the Bacillus globigii sample. For the Burkholderia, the nonmatching fraction is ~30% higher than Bacillus globigii, and this additional biological “noise” is reflected in an even higher value. While clearly our hypothesis test is sensitive to this population structure, it does tell us something useful about the sample and again highlights the importance for standardization of growth conditions, and potentially multiple calibrants to represent the population evolution tendencies of difference classes of organisms, as well as to capture potential sequence-dependent (i.e., high GC content) error processes. We note that the higher population diversity in the Burkholderia is also reflected in the number of SNPs called by our Pipeline B analysis.


Organism -score
nonmapping
Nonreference
base fraction
(mapped reads)
Mapped read fractionUnmapped reference basesSNP calls

B.mallei3.060.0250.94166,439431
B.pseudomallei2.870.0270.98859365
B.globigii*0.020.0180.9900

The values for Burkholderia are reflective of the higher fraction of non-reference-matching base calls in these samples and is indicative of greater population diversity compared to the Bg calibrant sample. The larger unmapped base counts along the reference genomes (column 5) for Bm are due to insertion elements that are highly mobile within these genomes and promote re-arrangements. BFAST default parameters for assigning candidate locations to reads when there is a high multiplicity of candidate alignments across the reference genome resulted in these gaps. Representative mapping statistics: 10,000,000 Illumina reads, 100 bp. *Calibrant.

The values are probability scores, calculated for observed mean signature coverage from sequence reads. value (statistic) are derived by subtracting actual measured from the interpolated value, representing this difference in units of standard deviations. values < ~2.0 given probability values <~0.01. To illustrate distribution of values obtained for unfiltered reads at mean signature coverage up to 150x, we plot values measured using within run calibration and external calibration.

The value represents the expected value from true match to a reference. The measured value (the hypothesis test) is the measurement on actual run data. To achieve accurate genome identification from populations of reads against reference database genomes using the calibration, we compute mean signature coverage for the reads against the reference, then interpolate from the calibration to estimate the predicted for an observed mean signature coverage for a true match.

One final and interesting aspect of the alignment/mapping pipeline analysis is seen in the fourth and fifth columns of Table 2. While the Burkholderia had greater genetic diversity than Bacillus globigii, B. mallei samples had lower fractions of successfully mapped reads, and a high fraction ~3% of unmapped based positions (gaps) in the reference genome. This can be explained by the fact that B. mallei genomes are unusually rich in identical insertion sequences which are mobile within the genome and promote genomic rearrangements. A deeper analysis of the gaps in the reference in our case shows they exist due to algorithmic choices in use of the BFAST tool for aligning reads that have a large number of candidate locations (assign to all locations versus assigning them at random) which results in underrepresented regions [11]. This represents another algorithmic caveat when dealing with genomes with high degrees of repetition.

When trying to detect SNPs that are useful for distinguishing related strains, it is valuable to compare a broad sampling of sequence information from subspecies being studied. Herein, we sequenced twenty-eight samples from the passage experiments. Tables 3 and 4 illustrate sequence variation detected when clones from the final passage where compared to their lineage progenitors. These clones represent the diversity of the genetic populations in the original source vial. Several clones from each of the lineages were also sequenced and compared to themselves and across the bulk population. For example, samples 8_1_2 and 8_1_3 have identical genotypes for chromosome 1 as the reference (as expected), but there are also clones that have variation from the reference. This indicates the direction of diversity that specific lineages are moving in. Some SNPs are conserved across the population while others are unique. Characterization of populations is essential for attribution.


Genome positionReference0_10_20_30_40_50_80_100_110_128_1_18_1_28_1_38_1_48_1_58_1_68_3_18_5_18_6_18_8_18_9_18_10_18_11_18_12_1

27694CAA
30272GAC
198367CTTAT
208896GC
208898GC
208901GA
209253GAA
238223GAA
238226TAAA
238248GA
254143C
260159CG
309954GC
387701TG
417834CT
417940TA
439654CAAGG
468747GACC
473445TG
502149T
522021CA
535552GC
575316CA
575340TA
636702TG
637354CA
637366TCC
719754GA
748773TC
748778GA
752001GA
755855GC
827221GAAA
830204C
840516CT
846684CTT
856204GA
856229GTT
867860TG
879922GA
908149CGG
918155GA
918163GA


Genome positionReference0_10_20_30_40_50_80_100_110_128_1_18_1_28_1_38_1_48_1_58_1_68_3_18_5_18_6_18_8_18_9_18_10_18_11_18_12_1

16302CAA
63867TCC
112549AG
112589AG
125529TCC
153871CAA
255127CTTTTTT
257976A
289329T
334349GAA
371920GAA
401983CTT
429628GA
438505AGG
446580T
479883TG
482145GA
563692AGG
568009GAA
656370CA
673008GTCCTT
767675CTT
767835TCC
769495CA
770544CA
790672CAA
793808GAA
794151GAA
794210GA
794352GT
794608TG
794942GAA
821535GA
859546AGG
859585GTT

We compared sequence information for samples across all twelve lineages that were derived from a common ancestor. For example, comparison of 8_10_1 to its progenitor shows that the genotypes are the same, as is the case for sample 8_12_1. Other samples have variants that are not in their progenitor, illustrating the complexity of sequence comparison of populations from a single source. We reduced these complexities to be able to derive a process to compare samples and their populations. The comparison is two-fold: (1) comprehensive analysis across the entire genome, and (2) population-wide comparison of clones to characterize the diversity of each sample. It is on account of sample’s diversity that making matches of samples based on population characterization improves accuracy of attribution.

4. Conclusion

Leveraging whole genome sequencing analysis strategies, entire genetic content of samples (populations) can be detected and identified as major and minor components. We have demonstrated effective analysis of lineage populations. In addition, we mapped the direction of mutation and compared taxonomic relationships and assembled genomes even when there were minor differences between genomes from the same source (see Figure 5). Using biothreat agents, Burkholderia pseudomallei and Burkholderia mallei, we cultured isolates, following mutations during passage, sequenced members of each population, built their individual genomes, and phylogenetically compared their relationships. These populations were measured and their diversity mapped with passage, which in turn enables traceability and attribution to a single source (Figure 3). We identified variation of these sample populations, indicating the unique mutations that are the DNA fingerprints of the sample profile and determined the entire population structure in the sample. We made comparisons against the genomes and direct comparison of sequence content between samples.

The diversity between passaged samples tracks with the lineages and in some cases there is overlap in genetic distance between members of the populations in different lineages, as expected (Figure 4). Overall, the pattern of diversity is spreading out from each lineage as a sphere of diversity. Hence, for forensics, the passaged material’s population is dependent on the source’s original content and the sequenced material is attributed back to the source. While individual members of the population are diverging and more distant phylogenetically, they are part of the sample’s profile and match back to the source. Comprehensive analysis across the populations points the results (passaged samples as evidence) of each sample (even though sequence reveals that each is not 100% identical match to other members of the population) back to the single source.

We applied the -value hypothesis testing approach to “detect” the specific threat agents we sequenced in multiple runs using calibration and quality control measures. In each case, a reference sequence with the same strain name as the sequenced organism was available in the Genbank database: Burkholderia mallei strain China 7 (ATCC 23744) and Burkholderia pseudomallei strain 668. The sequences were from a single colony, which corresponds to a single constituent within the parent population (source). Generally, during growth, this bacterium will accrue new random (and fitness-selected) mutations and rearrangements that result in consensus and population differences in the sequence content with respect to the reference genome.

5. Mapping Reads

Table 2 illustrates alignment-based mapping statistics for threat agents and our calibrant for comparison to nonalignment values. The higher values for Burkholderia are reflective of the higher fraction of non-reference-matching base calls in these samples, and is indicative of greater population diversity compared to calibration. Burkholderia had greater SNP calls compared to Y. pestis. The larger unmapped base counts along the reference genomes for Y. pestis and B. mallei are due to insertion elements that are highly mobile within these genomes and promote rearrangements. In addition to sample diversity, there are sequencing errors and artifacts to account for. Unlike other groups that ignore sequencing artifacts, we account for system errors to assign candidate locations to reads when there is a high multiplicity of candidate alignments.

Pipeline B enabled appropriate relationship evaluation of the populations because these samples all contain a continuum of variants. In such circumstances, SNP calling is not terribly meaningful way to approach the problem, and methods like SOAP have inherent algorithm bias that reduces their use for relationship analyses, because SNPs which occur close to other SNPs produce low confidence calls. Our goal was to evaluate the continuum of variation (major SNPs, minor SNPs, and background noise).

There are several advantages of this approach. The calibrant tools triage the sequence data, like a funnel, to improve the processing of accurate confident data instead of using computing time on background that masks interpretation of the populations. The system error generated that noise, but it is dependent on starting sequence input; hence, variable from sample to sample, this means that millions of sequence reads from Illumina data cannot be compared from sample to sample without the calibrant.

It is clear that direct population-sequencing characterizes entire genomes as biomarkers for sample profile comparisons. However, it is important to understand the mechanisms behind probabilistic matching approaches. Herein, for example, in this case, the SNP calls, for pipeline A, are above the confidence criteria, but not called because the high number of SNPs in a small window of sequence. Hence, pipeline B provides accurate sequence information for tracking populations and matching samples to source.

If more conservative calls are made, then some SNP calls by only one algorithm could be assigned a lower confidence score. It is not so much a question as to which algorithm is more accurate as a question of the level of confidence for the specified conditions. Users need to understand the criteria they are satisfied with.

Pipeline B processed reads to establish the edge of the “background” noise, projecting a threshold to set a false call rate against this background of 0.1/genome analyzed, and then called SNPs above that threshold. Thus, for comparison purposes, how each user will choose to treat the population will determine whether the analyses will coincide. Analysis of the population contained in the sample is the key to forensic attribution. Comprehensive analysis of the genetic constituents establishes the sample profile. Profiles can be compared to determine the relatedness of the populations in the samples, not just one or two selected targets. Comprehensive analysis also strengthens the probability of the match and increases the confidence to include and exclude samples as matches.

Overall, direct sequence analysis of populations, as a biomarker, reveals the variation of genomes. Our experimental design increased homogeneity produced by single cell bottlenecks allowed a controlled analysis of the data. Many derivatives of experimental design could be extended from our work to further characterize the extent of genome variation within populations and strategies to employ to use sequencing information across entire genomes as biomarkers. With the increasing availability of complete genome sequence data from multiple microbial pathogens, comparative genomics has recently emerged as a powerful tool to understand the basic molecular properties of pathogens. Hence, population-sequencing can be used as a direct method to identify disease agents and genomes, rendering the entire genetic content, variation and diversity as the biomarker(s) for detection. Our study confirms utility of alignment-free and alignment-based approaches in defining the presence and significance of genomic variation in populations of B. pseudomallei and B. mallei. Further studies are required to determine the full extent of variability and their relationship to biological fitness in the environment and to disease pathogenesis in the human host.

Finally, sequencing analysis is highly sensitive in tracing genomic differences among isolates. Direct use of sequencing for characterization of populations of genomes present in samples provides insights into the number and types of microorganisms present. While biomarkers targeted on detection of a disease agent at the genus or species levels may indicate a threat is present, comprehensive analysis of variants, near-neighbors, and the community structure will improve the accuracy of disease detection and identification of the causal agent adding important information about disease context and community. Using DNA elements across the entire genome represents a better biomarker than PCR probes, which target limited information. Population-sequencing identifies the disease agent, provides insights on variation, and captures information about more than just a single target, indicating multiple viral and bacterial pathogens and commensals contained in samples.

Acknowledgments

This study was funded by the Department of Homeland Security contract Whole Genome Approach to Microbial Forensics (WGAMF) HSHQDC-10-C-00140 and IAA no. W911SR-10-C-0026 Bioagent Early Warning Detector, (BEWD), RDT&E, Chemical and Biological Defense Program, Defense Wide, Line 14, PE 0602384BP. Samples were prepared and handled at the CUBRC BSL facility and cultured under standard procedures. DNA from serial passages of biothreat agents was extracted and sequenced. Sequencing was conducted at ECBC as per Illumina recommended protocols.

References

  1. C. Trapnell and S. L. Salzberg, “How to map billions of short reads onto genomes,” Nature Biotechnology, vol. 27, no. 5, pp. 455–457, 2009. View at: Publisher Site | Google Scholar
  2. H. Nagasaki, T. Mochizuki, Y. Kodama et al., “DDBJ read annotation pipeline: a cloud computing-based pipeline for high-throughput analysis of next-generation sequencing data,” DNA Research, vol. 20, no. 4, pp. 383–390, 2013. View at: Publisher Site | Google Scholar
  3. T. Camerlengo, H. G. Ozer, R. Onti-Srinivasan et al., “From sequencer to supercomputer: an automatic pipeline for managing and processing next generation sequencing data,” AMIA Summits on Translational Science Proceedings, vol. 12, pp. 1–10, 2012. View at: Google Scholar
  4. G. A. Pavlopoulos, A. Oulas, E. Iacucci et al., “Unraveling genomic variation from next generation sequencing data,” BioData Mining, vol. 6, no. 1, article 13, 2013. View at: Google Scholar
  5. J. P. Jakupciak and R. R. Colwell, “Biological agent detection technologies,” Molecular Ecology Resources, vol. 9, supplement 1, pp. 51–57, 2009. View at: Publisher Site | Google Scholar
  6. K. A. Crandall, C. R. Kelsey, H. Imamichi, H. C. Lane, and N. P. Salzman, “Parallel evolution of drug resistance in HIV: failure of nonsynonymous/synonymous substitution rate ratio to detect selection,” Molecular Biology and Evolution, vol. 16, no. 3, pp. 372–382, 1999. View at: Google Scholar
  7. J. E. Cooper and E. J. Feil, “Multilocus sequence typing—what is resolved?” Trends in Microbiology, vol. 12, no. 8, pp. 373–377, 2004. View at: Publisher Site | Google Scholar
  8. O. E. Francis, M. Bendall, S. Manimaran et al., “Pathoscope: species identification and strain attribution with unassembled sequencing data,” Genome Research, vol. 23, no. 10, pp. 1721–1729, 2013. View at: Publisher Site | Google Scholar
  9. M. Wang, Y. Ye, and T. Haixu, “A de Bruijn graph approach to the quantification of closely-related genomes in a microbial community,” Journal of Computational Biology, vol. 19, pp. 814–825, 2012. View at: Google Scholar
  10. J. P. Jakupciak, “Population-sequencing as a biomarker for sample characterization,” Journal of Biomarkers. In press. View at: Google Scholar
  11. J. P. Jakupciak, J. M. Wells, J. S. Lin, and A. B. Feldman, “Population analysis of bacterial samples for individual identification in forensics application,” Journal of Datamining in Genomics & Proteomics, vol. 4, article 138, 2013. View at: Publisher Site | Google Scholar
  12. M. T. G. Holden, R. W. Titball, S. J. Peacock et al., “Genomic plasticity of the causative agent of melioidosis, Burkholderia pseudomallei,” Proceedings of the National Academy of Sciences of the United States of America, vol. 101, no. 39, pp. 14240–14245, 2004. View at: Publisher Site | Google Scholar
  13. K. T. Konstantinidis, A. Ramette, and J. M. Tiedje, “The bacterial species definition in the genomic era,” Philosophical Transactions of the Royal Society B, vol. 361, no. 1475, pp. 1929–1940, 2006. View at: Publisher Site | Google Scholar
  14. B. J. Currie, A. Haslem, T. Pearson et al., “Identification of melioidosis outbreak by multilocus variable number tandem repeat analysis,” Emerging Infectious Diseases, vol. 15, no. 2, pp. 169–174, 2009. View at: Publisher Site | Google Scholar
  15. K. H. Chua, K. H. See, K. L. Thong, and S. D. Puthucheary, “DNA fingerprinting of human isolates of Burkholderia pseudomallei from different geographical regions of Malaysia,” Tropical Biomedicine, vol. 27, no. 3, pp. 517–524, 2010. View at: Google Scholar
  16. C.-S. Chiou, “Editorial: multilocus variable-number tandem repeat analysis as a molecular tool for subtyping and phylogenetic analysis of bacterial pathogens,” Expert Review of Molecular Diagnostics, vol. 10, no. 1, pp. 5–7, 2010. View at: Publisher Site | Google Scholar
  17. E. P. Price, H. M. Hornstra, D. Limmathurotsakul et al., “Within-host evolution of Burkholderia pseudomallei in four cases of acute melioidosis,” PLoS Pathogens, vol. 6, no. 1, Article ID e1000725, 2010. View at: Publisher Site | Google Scholar
  18. C. M. Ronning, L. Losada, L. Brinkac et al., “Genetic and phenotypic diversity in Burkholderia: contributions by prophage and phage-like elements,” BMC Microbiology, vol. 10, pp. 202–208, 2010. View at: Publisher Site | Google Scholar
  19. M. A. Schell, R. L. Ulrich, W. J. Ribot et al., “Type VI secretion is a major virulence determinant in Burkholdria mallei,” Molecular Microbiology, vol. 64, no. 6, pp. 1466–1485, 2007. View at: Google Scholar
  20. A. Deeraksa, O. Qazi, G. C. Whitlock et al., “Development of a non-living vaccine against Burkholderia mallei,” The Journal of Immunology, vol. 182, article 129. 2, 2009. View at: Google Scholar
  21. D. A. B. Dance, “Melioidosis: the tip of the iceberg?” Clinical Microbiology Reviews, vol. 4, no. 1, pp. 52–60, 1991. View at: Google Scholar
  22. E. Yabuuchi and M. Arakawa, “Burkholderia pseudomallei and melioidosis: be aware in temperate area,” Microbiology and Immunology, vol. 37, no. 11, pp. 823–836, 1993. View at: Google Scholar
  23. L. D. Rotz, A. S. Khan, S. R. Lillibridge, S. M. Ostroff, and J. M. Hughes, “Public health assessment of potential biological terrorism agents,” Emerging Infectious Diseases, vol. 8, no. 2, pp. 225–230, 2002. View at: Google Scholar
  24. D. A. Rasko, P. L. Worsham, T. G. Abshire et al., “Bacillus anthracis comparative genome analysis in support of the Amerithrax investigation,” Proceedings of the National Academy of Sciences of the United States of America, vol. 108, no. 12, pp. 5027–5032, 2011. View at: Publisher Site | Google Scholar
  25. M. J. Struelens, Y. De Gheldre, and A. Deplano, “Comparative and library epidemiological typing systems: outbreak investigations versus surveillance systems,” Infection Control and Hospital Epidemiology, vol. 19, no. 8, pp. 565–569, 1998. View at: Google Scholar
  26. S. N. Gardner, C. J. Jaing, K. S. McLoughlin, and T. R. Slezak, “A microbial detection array (MDA) for viral and bacterial detection,” BMC Genomics, vol. 11, no. 1, article 668, 2010. View at: Publisher Site | Google Scholar
  27. P. H. M. Savelkoul, H. J. M. Aarts, J. De Haas et al., “Amplified-fragment length polymorphism analysis: the state of an art,” Journal of Clinical Microbiology, vol. 37, no. 10, pp. 3083–3091, 1999. View at: Google Scholar
  28. P. Janssen, R. Coopman, G. Huys et al., “Evaluation of the DNA fingerprinting method AFLP as a new tool in bacterial taxonomy,” Microbiology, vol. 142, part 7, pp. 1881–1893, 1996. View at: Google Scholar
  29. B.-A. Lindstedt, E. Heir, T. Vardund, and G. Kapperud, “A variation of the amplified-fragment length polymorphism (AFLP) technique using three restriction endonucleases, and assessment of the enzyme combination BglII-MfeI for AFLP analysis of Salmonella enterica subsp. enterica isolates,” FEMS Microbiology Letters, vol. 189, no. 1, pp. 19–24, 2000. View at: Publisher Site | Google Scholar
  30. Y. Graser, I. Klare, E. Halle et al., “Epidemiological study of an Acinetobacter baumannii outbreak by using polymerase chain reaction fingerprinting,” Journal of Clinical Microbiology, vol. 31, no. 9, pp. 2417–2420, 1993. View at: Google Scholar
  31. B. Budowle, M. D. Johnson, C. M. Fraser, T. J. Leighton, R. S. Murch, and R. Chakraborty, “Genetic analysis and attribution of microbial forensics evidence,” Critical Reviews in Microbiology, vol. 31, no. 4, pp. 233–254, 2005. View at: Publisher Site | Google Scholar
  32. M. A. Poritz, A. J. Blaschke, C. L. Byington et al., “FilmArray, an automated nested multiplex PCR system for multi-pathogen detection: development and application to respiratory tract infection,” PLoS ONE, vol. 6, no. 10, Article ID e26047, 2011. View at: Publisher Site | Google Scholar
  33. J. K. Stone, M. Mayo, S. A. Grasso et al., “Detection of Burkholderia pseudomallei O-antigen serotypes in near-neighbor species,” BMC Microbiology, vol. 12, article 250, 2012. View at: Google Scholar
  34. I. Vandenbroucke, H. van Marck, P. Verhasselt et al., “Minor variant detection in amplicons using 454 massive parallel pyrosequencing: experiences and considerations for successful applications,” BioTechniques, vol. 51, no. 3, pp. 167–177, 2011. View at: Publisher Site | Google Scholar
  35. J. W. Davey, P. A. Hohenlohe, P. D. Etter, J. Q. Boone, J. M. Catchen, and M. L. Blaxter, “Genome-wide genetic marker discovery and genotyping using next-generation sequencing,” Nature Reviews Genetics, vol. 12, no. 7, pp. 499–510, 2011. View at: Publisher Site | Google Scholar
  36. C.-C. Ho, C. C. Y. Lau, P. Martelli et al., “Novel pan-genomic analysis approach in target selection for multiplex PCR identification and detection of Burkholderia pseudomallei, Burkholderia thailandensis, and Burkholderia cepacia complex species: a proof-of-concept study,” Journal of Clinical Microbiology, vol. 49, no. 3, pp. 814–821, 2011. View at: Publisher Site | Google Scholar
  37. S. A. Boers, W. A. van der Reijden, and R. Jansen, “High-throughput multilocus sequence typing: bringing molecular typing to the next level,” PLoS ONE, vol. 7, Article ID e39630, 2012. View at: Google Scholar
  38. M. V. Larsen, S. Cosentino, S. Rasmussen et al., “Multilocus sequence typing of total-genome-sequenced bacteria,” Journal of Clinical Microbiology, vol. 50, no. 4, pp. 1355–1361, 2012. View at: Publisher Site | Google Scholar
  39. M. Inouye, T. C. Conway, J. Zobel, and K. E. Holt, “Short Read Sequence Typing (SRST): multi-locus sequence types from short reads,” BMC Genomics, vol. 13, article 338, 2012. View at: Google Scholar
  40. S. J. Shallom, H. Tae, L. Sarmento et al., “Comparison of genome diversity of Brucella spp. field isolates using Universal Bio-signature Detection Array and whole genome sequencing reveals limitations of current diagnostic methods,” Gene, vol. 509, no. 1, pp. 142–148, 2012. View at: Google Scholar
  41. K. Lagesen, D. W. Ussery, and T. M. Wassenaar, “Genome update: the 1000th genome—a cautionary tale,” Microbiology, vol. 156, no. 3, pp. 603–608, 2010. View at: Publisher Site | Google Scholar
  42. T. C. Glenn, “Field guide to next-generation DNA sequencers,” Molecular Ecology Resources, vol. 11, no. 5, pp. 759–769, 2011. View at: Publisher Site | Google Scholar
  43. M. L. Metzker, “Sequencing technologies the next generation,” Nature Reviews Genetics, vol. 11, no. 1, pp. 31–46, 2010. View at: Publisher Site | Google Scholar
  44. S. Pabinger, A. Dander, M. Fischer et al., “A survey of tools for variant analysis of nest generation genome sequencing data,” Briefings in Bioinformatics, 2013. View at: Publisher Site | Google Scholar
  45. M. Eppinger, P. L. Worsham, M. P. Nikolich et al., “Genome sequence of the deep-rooted Yersinia pestis strain Angola reveals new insights into the evolution and pangenome of the plague bacterium,” Journal of Bacteriology, vol. 192, no. 6, pp. 1685–1699, 2010. View at: Google Scholar

Copyright © 2013 John P. Jakupciak et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


More related articles

 PDF Download Citation Citation
 Download other formatsMore
 Order printed copiesOrder
Views1710
Downloads857
Citations

Related articles

Article of the Year Award: Outstanding research contributions of 2020, as selected by our Chief Editors. Read the winning articles.