Ultra-deep mutant spectrum profiling: improving sequencing accuracy using overlapping read pairs
© Chen-Harris et al.; licensee BioMed Central Ltd. 2013
Received: 17 October 2012
Accepted: 6 February 2013
Published: 12 February 2013
High throughput sequencing is beginning to make a transformative impact in the area of viral evolution. Deep sequencing has the potential to reveal the mutant spectrum within a viral sample at high resolution, thus enabling the close examination of viral mutational dynamics both within- and between-hosts. The challenge however, is to accurately model the errors in the sequencing data and differentiate real viral mutations, particularly those that exist at low frequencies, from sequencing errors.
We demonstrate that overlapping read pairs (ORP) -- generated by combining short fragment sequencing libraries and longer sequencing reads -- significantly reduce sequencing error rates and improve rare variant detection accuracy. Using this sequencing protocol and an error model optimized for variant detection, we are able to capture a large number of genetic mutations present within a viral population at ultra-low frequency levels (<0.05%).
Our rare variant detection strategies have important implications beyond viral evolution and can be applied to any basic and clinical research area that requires the identification of rare mutations.
KeywordsQuasispecies Viral evolution DNA mutational analysis High-throughput sequencing Diagnostics Biomarker Rare mutations Sequencing error correction Overlapping read pairs
Viruses with RNA genomes replicate with extremely high mutation rates because their RNA polymerases lack the proofreading ability of DNA polymerases. With a mutation rate of ~1 error per 10,000 nucleotides copied, a point mutation is introduced nearly every time a single RNA virus replicates . Any given viral sample extracted from a host contains a spectrum of related genotypes, referred to as a quasispecies, whose ability to rapidly evolve underlies viral virulence, vaccine resistance and host-jumping . Understanding the mutational dynamics of RNA viruses is key to our understanding of viral disease progression, transmission and the development of antiviral therapeutics.
Considerable progress has been made recently using deep sequencing to characterize the mutant spectra in several human RNA viral pathogens: human immunodeficiency virus (HIV) [3–6], hepatitis C virus (HCV) [7–9] and influenza [10, 11]. In particular, deep sequencing has been used to identify medically relevant drug-resistant rare variants that impact anti-retroviral drug treatment outcomes [11–15]. Many of the earliest efforts have targeted specific hyper variable genomic regions using 454 sequencing; more recently published studies have begun to target more of the genome with high depths using the greater sequencing output of Illumina technologies [16, 17]. Recently, the software tool for managing deep sequencing data, Segminator II, was introduced and used to compare performance of Illumina and 454 deep sequencing of influenza . However, less emphasis was given to evaluating a general subconsensus base calling procedure and the impact of PCR amplification was not considered. Two other available packages that address PCR errors and known sequencing error modes, AmpliconNoise  and RC454 , are designed specifically for 454 pyrosequencing data. In a recent Illumina deep sequencing study of foot-and-mouth disease virus samples, Wright et al.  counted evidence for between 1,434 and 2,622 rare variants present in their samples. Their approach relied on an estimated error rate from the sequencer without the use of sequencing controls and included sequencing each sample twice to correct for sequencing error, which would present practical problems for sequencing larger numbers of samples collected from an outbreak. Moreover, recent work has indicated the presence of non-uniform error rates in Illumina sequence data in particular, and highlights the ongoing challenge of correctly separating the true mutant spectra from sequencing related errors [15, 22–24].
To date, few investigations have applied high throughput sequencing on viruses that naturally circulate in animal hosts. Given the potential for RNA viruses that circulate in a non-human host reservoir to infect new host types, including humans , it is important to study viral evolution at the highest resolution for measuring genetic change. Characterizing these viruses pose a particular challenge since the starting material can be too small (or rare) to directly sequence without PCR amplification or growth in cell culture, which can introduce new errors to the measurement process . As a pilot study for a series of viral evolution studies, three viral samples collected from naturally infected hosts--two fox rabies brain tissue samples and one bovine coronavirus (BCV) nasal sample–were sequenced using Illumina paired-end read technology at ultra-deep coverage (> 300,000x raw reads). Two plasmid clones containing 1 kb region of the rabies and BCV genomes served as error controls in the study. Error rates seen in plasmids provided a best estimate of the combined PCR and sequencing error rate for the natural samples .
Overlapping paired-end reads, generated using short sequencing fragment libraries combined with long read lengths, have been recently used to improve Illumina paired-end read assemblies in software packages such as FLASH  and PANDAseq , in which overlapping regions of the read pairs serve to extend read lengths and reduce sequencing errors. Overlapping read pairs have not been previously applied to mutant spectra profiling to improve the sensitivity of variant detection. We demonstrate the novel use of mismatch rates in the overlapping read pairs (ORP) to provide an unbiased assessment of the sequencer-derived quality scores when selecting a read filtering threshold, as well as to estimate position-dependent sequencing errors without relying on a clonal control. Our methods identified a high variation in error rates between sequencers and indicated the difficulty of relying on traditional sequencing approaches for rare variant characterization. Moreover, we demonstrate that PCR amplification can become the dominant source of error over the sequencer’s error even when using a high fidelity polymerase. Among the three natural viral samples we sequenced, we identified up to 2,133 rare variants within a single sample with an in-host population frequency as rare as 0.0096%. Given the majority of the variants we discovered occurred at the ultra-rare level (< 0.1%), we show that careful error control and estimation using ORP can reveal a deep and rich mutant spectrum. Our results demonstrate a practical sequencing and computational analytic approach to studying viral evolution with an unprecedented level of genetic resolution.
Results and discussion
ORPs reduce error rates and provide benchmarks for quality scores
Sequence coverage for the three natural RNA viral samples and the two plasmid control samples
No. bases sequenced
Raw/single read coverage
ORP (Q≥30) coverage
Singleton ends coverage
The long overlapping regions in the read pairs offered several important practical benefits to improving the quality of the reads. First, they served as a mechanism of error checking, as each read pair came from the same template and should therefore be perfectly complementary. Base calls that do not match in the forward and reverse strands are automatically identified as sequencing errors. Second, although the mismatched bases were excluded from data analysis, they provided an empirical estimate for single-read sequencing error rates. Third, it identified “problematic loci” on the genome where large fractions of the ORPs are mismatched. A high fraction of ORP mismatches at a particular locus would indicate that the locus was a site with high probability of erroneous nucleotide incorporation and hence suggest that a more stringent criterion should be considered when making variant calls at the locus.
Overall mismatch rates in overlapping paired-end reads for raw read pairs, read pairs that have quality scores Q10, Q≥20, Q≥30 and Q≥35
Average of 5 samples
Rabies control –repeat run
The Q-score associated with each base-call is derived based on an aggregate of sequencer metrics and sample characteristics measured during each sequencing run. Q-scores of 10, 20, 30 and 35 correspond to error rates of 0.1, 0.01, 0.001, 0.0003, respectively. The mean mismatch rates among the raw, Q10, Q20, Q30 and Q35 ORPs were 6.7×10-3, 8.8×10-4, 1.2×10-4, 4.0×10-6, 2.5×10-7, respectively (Table 2, average of 5 samples), far lower than what their Illumina Q-scores suggest. However, when the rabies control plasmid was sequenced a second time, the resulting ORP mismatch rate profile was quite different. Though the mismatch rates were comparable to the first run in the raw, unfiltered ORP reads, they were significantly higher in the second run when comparing the same Q thresholds (Table 2). At Q30, ORP mismatch rates in the first run were two orders of magnitude lower than the rabies control repeat run. Given the same model of Next Generation Sequencing instrument (Illumina GA IIx) was used in both sequencing runs, the most parsimoneous explanation for the difference in ORP mismatch rates is the difference in the Q-score calibration of the two instruments. This underscores the utility of using ORP to recover an empirical sequencing error rate and minimize technical artifacts introduced by the sequencer.
The trade-off for enhanced accuracy is the reduction in coverage. Since the two overlapping reads represent redundant information from the same amplicon, the true depth of coverage is the number of read pairs instead of number of single reads. For example, in the BCV control sample, the Q-score filtering process removes roughly 11%, 22% and 55% of the raw matching ORPs at each base at Q10, Q20 and Q30, respectively (see Additional file 1: Figure S1).
ORPs help distinguish PCR error rates from sequencing error rates
Summary of per base-call error rates in the two control sequences
Q10 mean error rate
Q10 max error rate
Q30 mean error rate
Q30 max error rate
Our variant detection model uses a position-dependent error rate that takes the mismatch rate found at each base into consideration (Methods). Before applying this model on the natural samples, we applied it to the control data and determined that the false positive rates associated with 3 error rates, 5×10-5, 1×10-4 and 5×10-4 were 2.45%, 0.75% and 0%, respectively (Additional file 1: Table S2). This suggests that the mean Q30 ORP error rate of ~5×10-5 estimated from the control data (Table 3) was not conservative enough to eliminate all false positive variant calls, possibly due to PCR errors. Based on these observations, we made variant calls in the three natural viral samples using both 5×10-5 and a more conservative error rate of 5×10-4, where the former represented the mean overall error rate and the latter approximated the PCR error rate.
Rare variants found at 10-13% of the genomes in the natural viral samples
Summary of coverage, total number of candidate variants (all polymorphisms) and number variants called by the variant detection model in Q30 ORP for the 3 natural viral samples
Mean ORP coverage
# candidate variant calls at Q≥30
# variants called at error rate 5×10-5
# variants called at error rate 5×10-4
FDR for error rate 5×10-5
FDR for error rate 5×10-4
Recovering the bulk of these ‘ultra-rare’ viral mutations, however, requires considerable extra effort – either through reduced error rate or increased coverage. Since for most viral evolution study designs it may not be practical or necessary to sequence viral samples at the coverage level used in this paper, we estimated the theoretical coverage required to achieve specific variant detection sensitivity under three error rates: 5×10-5, 1×10-4, and 5×10-4. Predictions are based on a binomial error model with a fixed Bonferroni correction factor (assuming 11,000 candidate variants detected in every case) and shown in Figure 5B. For instance, at 20,000x coverage (matching ORP), the maximum sensitivity, or the rarest variants that can be detected at these three error rates have population frequencies of 0.05%, 0.065% and 0.145%, respectively, with the lowest error rate (5×10-5) being the most sensitive and the highest error rate (5×10-4) being the least sensitive.
As shown in Figure 4, accuracy of Q30 single reads is comparable to that of Q20 ORPs. In the scenario of 20,000x ORP coverage, if non-overlapping read pairs had been generated during sequencing, in theory, the number of Q30 single reads could potentially double the coverage to 40,000x. At the error rate of 5×10-5, the maximum sensitivity for variant detection in these Q30 single reads would be 0.03% instead of 0.05%. The cost of this increased coverage by generating non-overlapping read pairs is losing the benefit of context-specific error estimation and correction afforded by the overlapping read pairs. The accuracy comparison between Q20 ORPs and Q30 single reads observed in our data set was only established because ORPs were used. Any accuracy equivalence between single reads and ORP has to be carefully re-established for the specific sequencer being used.
In order to effectively use ultra-deep sequencing to study rare members of a viral population, it is critical to accurately model errors in the sequencing data. We have developed a protocol to evaluate and control sequencing error in multiple ways. First, plasmid clones were sequenced along with viral samples of interest so that error rates in the PCR and sequencing process could be empirically derived. Second, taking advantage of the Illumina paired-end technology, overlapping paired-end reads were generated to improve read accuracy. Third, mismatch rates in the ORP and error rates in the control plasmids were examined in association with quality scores from the sequencers so an optimal Q-score threshold could be selected for read filtering. Mismatch rates were also incorporated in the variant detection model to dynamically adjust error rates based on local sequencing errors. Applying this model on the plasmid control data before the natural samples further gave empirical assessment of false positive rates in the data for a given error rate.
Sequencing errors were directly estimated from mismatch rates in the ORP. We found that these Q-score filtered ORPs had sequencing errors far below what their Q-scores suggested. Using mismatch rates in the ORP, we demonstrated considerable variability of quality metrics on two Illumina GA IIx sequencers (Table 2). This variability may stem from differences in the Q-score calibration process on the sequencers. Together, these findings suggest that Q-scores by themselves are not reliable measures of sequencing accuracy. Mismatch rates in ORPs, however, provide unbiased estimates on sequencing error and can be used to select Q-scores for read filtering.
Among the challenges of correctly separating the true mutations from sequencing related errors is the presence of the non-uniform error rates in the sequencing data [15, 20–22]. We have shown that even at Q30, ORP mismatch rates have a small but significant distribution (Figure 2). Mismatch rates in the ORPs offer locus-specific information on error rates and thus can improve variant call accuracy. Besides the Illumina paired-end technology, repeated measurements on the same read fragments can also be generated using other sequencing platforms such as the PacBio RS from PacificBiosciences . As with techniques developed in this work, future new data generation methods that support high-throughput repeated interrogation of the same insert fragment and informatics techniques that exploit this feature can also be used in combination to substantially reduce the possibility of error.
ORP has potentially greater benefits in the application of direct sequencing without PCR. Our results show that with careful quality control, the accuracy of PCR-amplicon sequencing will be limited by PCR rather than sequencing errors. While it is often necessary to use RT-PCR to amplify viral RNA from host material, the exponential nature of the PCR reaction combined with issues such as primer bias can skew the variant frequencies (e.g. toward laboratory-derived reference strains) and even generate incorrect consensus sequences in some regions of the genome. This highlights the importance of pursuing alternative, direct sequencing technologies. Until recently, it has not been practical to sequence viral samples such as ours without PCR amplification. As the technology of direct sequencing improves and becomes available, methods that can reduce sequencing error, such as the use of ORP, will play a greater role in quality control.
The tradeoff for enhanced accuracy by using ORP and Q-score filtering is the reduction in coverage. Choosing to generate overlapping read pairs instead of non-overlapping read pairs can reduce the effective coverage by half. Another limitation of ORPs is the effective shortening of read length, which limits downstream analysis such as linking multiple sub-consensus mutations to a single haplotype. Nevertheless, currently the longest length of non-overlapping read pairs is still too short to span an entire gene or genome for the purpose of reconstructing subconsensus haplotypes.
Several alternative ultra-sensitive mutation detection approaches have been proposed recently for next generation sequencing [15, 31, 32], but they require significant sample preparation and may not be practical beyond targeting limited regions of a genome. Both Duplex Sequencing  and Safe-SeqS  require labeling the DNA fragment libraries with unique sequence tags (UID) prior to PCR amplification. Post-sequencing, mutations that occur in the majority of their uniquely tagged read families are identified as true variants. These methods successfully address the errors introduced during PCR and sequencing but are subject to the efficiency of the UID assignment. A significant fraction of the starting material is generally lost in the library prep procedures for Illumina sequencing due to poor adapter ligation efficiency and the requirement of multiple clean-up cycles. If a sample contains limited starting material for sequencing, as is often the case in viral or clinical samples, performing adapter ligation prior to PCR amplification will likely lead to poor representation of the sample. Furthermore, the additional UID assignment process adds to the complexity and cost of sample preparation. Thus while these methods present a possible approach for the future, their scalability to whole genome sequencing has not yet been demonstrated. Flaherty et al.  proposed an ultrasensitive mutation detection method for targeted resequencing using a position-specific error profile. They derive the position-specific error profile of a 700 bp region of the NA gene in H1N1 using a clone of the sample of interest. This approach while highly specific, also does not scale to whole genome analysis. In contrast to these methods, we suggest that the use of ORPs to derive error profiles is easily extendible to a genome of any size without the use of a cloned sample. Comparable fragment length and read length is the only requirement to generate ORPs.
This study for the first time provides a detailed analysis of the current standard single read approach versus overlapping read pairs in the application of mutant spectra profiling of viral samples. The results show there are compelling benefits to using overlapping read pairs, which lead us to favor their use whenever feasible. Yet the results also show that with careful filtering, single reads provide a viable option when PCR amplification is used in the sample preparation stage, and enough experience with the specific sequencer is available to eliminate the impact of unexpected changes in calibrated quality scores or other technical configuration changes. The advantage of ORPs is the dynamic context specific error estimation and error correction, which significantly reduces sequencing error and is more robust in the face of configuration changes across sequencing machines.
Sensitivity of viral variant detection is a function of error rate and coverage. We provided theoretical maximum sensitivity for a given coverage at three different error rates (Figure 5B). Consistent with these predictions, the majority of the variants detected in our data occurred at the frequencies of 0.01% to 0.1% for the error rate of 5×10-5. For our samples, the ‘ultra-rare’ mutants make up a majority of the sub-consensus population (Figure 5A). These ultra-rare mutants greatly increase the genetic diversity in the quasispecies and may play important roles in acute viral infectious diseases. While it remains an open question as to their ultimate importance for downstream applications, our results illustrate the significance of reduced error and increased coverage in recovering these rare mutations. Reliable rare variant detection and sequence error reduction is important for many research areas beyond virology. Our methods have applications in clinical diagnostics [15, 31], forensics  and DNA-based information storage system . Although high-throughput sequencing technologies have been limited in some applications due to their high error rates, repeated template sequencing presents a powerful approach for increasing the sequencing fidelity to a level that can generate highly sensitive detection assays.
This study was approved by the Institutional Animal Care and Use Committee at Lawrence Livermore National Laboratory (Protocol Number 2009–207). The rabies positive brain samples included in this study were not covered by IACUC 2009–207 as they were taken from residual archived diagnostic samples offered by the California Department of Public Health Laboratory, Richmond, CA and were originally collected and tested to inform public health decisions on administering anti-rabies vaccination.
The data described in this paper is available via anonymous ftp as reads, BAM format alignments, consensus sequence, nucleotide frequency profiles at: ftp://gdo144.ucllnl.org/pub/orpdat/.
Rabies: Two brain tissue samples obtained from grey foxes (Urocyon cinereoargenteus) displaying symptoms of rabies were collected in Humboldt County, CA in March 2009 and December 2009 and tested for rabies virus via RT-PCR  using a modified protocol that amplifies a portion of the N gene. Approximately 1 gram of tissue from each brain was sent to LLNL for analysis. RNA was extracted from the tissue sample using TRIzol® LS Reagent (Invitrogen, Carlsbad, CA) following the manufacturer’s protocol.
Bovine coronavirus (BCV): Nasal samples were collected from approximately 100 asymptomatic calves. Samples were collected using sterile polyester swabs, placed in 2-3 mL of Eagle’s Minimum Essential Medium (Gibco) and transported on ice back to the laboratory. Collected nasal swabs were vortexed in Eagle’s Minimum Essential Medium supplemented with 1% antibiotic-antimycotic solution (Gibco). The sample suspensions were clarified by centrifugation at 2000 x g for 30 minutes, filtered through a 0.22 μm filter and aliquots of about 500–1000 μl were stored at −80°C. RNA was extracted using TRIzol® LS Reagent following the manufacturer’s protocol. Samples containing BCV RNA were identified as described in Cho et al. .
Approximately 11 kb of the rabies virus genome and 12 kb of the BCV genome were amplified using reverse transcriptase (RT) PCR. PCR primer candidates were selected based on the combined results of the multiple sequence alignment and sequence searches. This technique is a modified version of the approach outlined in Slezak et al. . Three sets of degenerate PCR primers were tested for the amplification of each overlapping region of the rabies virus and BCV genomes using increments of 1.5-2.5 kb. For each region, the two primer sets that performed best were used to amplify cDNA obtained from the two fox rabies samples and one BCV sample. All primers used for amplification are given in Additional file 1.
RT-PCR and cloning
The rabies and BCV genomes were amplified using two-step RT-PCR using Superscript III RT reverse transcriptase kit and Platinum Pfx polymerase (Invitrogen), following manufacturer's instructions. Reverse transcription was performed using random hexamers the PCR conditions consisted of 94°C for 5 min, followed by 35 cycles of 94°C for 15 s, 54-60°C for 30 s, and 68°C for 2.5 min.
A 1 kb region of the rabies virus and BCV genome were amplified and each cloned into a plasmid vector. The inserts were generated by RT-PCR as described above using rabies and BCV polymerase primers: RVpolyF1 5’ CCCCTGACTCCTTATATCAAAACC, RVpolyR1 5’ GCGAGGTTGACTATTTGGTC, BCVpolyF2 5’ TTTGCAGACAAATTGGTGGA, and BCVpolyR2 5’GGCGTAAATTTCATCCTGCT. Poly 3’ A overhangs were added to the PCR products by incubating the products with Taq polymerase at 72°C for 10 min. TOPO TA Cloning Kit for Sequencing (Illumina) was used to clone the PCR products into One Shot® TOP10 cells (Invitrogen) as per manufacturer’s instructions. Sanger sequencing of the cloned controls was carried out by ELIM Biopharmaceuticals, Inc., Hayward CA. PCR products were prepared for Illumina sequencing using the QIAquick PCR Purification kit (Qiagen).
Sequencing of the three natural samples and the two control plasmids was carried out by Eureka Genomics (Hercules, CA) using an Illumina Genome Analyzer IIx. Each natural viral sample was sequenced in a separate lane of a single flow cell using paired-end reads on short genomic fragment inserts using read lengths of 112 bases. The clonal controls were mixed in a single sample with an approximate concentration ratio of 10:1 (rabies:BCV) and sequenced on a separate lane. Since the PCR primers could potentially introduce false mutations into the amplicon pool due to non-specific binding, primer regions were masked out for the downstream analysis. Table 1 summarizes the output generated in the sequencing runs.
To compare error rates between different sequencers, rabies control plasmid was sequenced a second time at Elim Biopharm (Hayward, CA) using Illumina Genome Analyzer IIx. A shorter fragment length was chosen with more complete overlap of the read pairs. The same amplicon pool was used for both sequencing runs.
Read mapping to reference
The open source software SHRiMP2 was used for read mapping. The tool was chosen for its high read mapping sensitivity  and its ability to map as many reads as possible in the face of individual errors within each read .
A consensus sequence was generated for each sample following an iterative comparative assembly procedure suggested by Willerth et al. . In this approach, an initial reference sequence was chosen, reads were mapped to the reference, then a new consensus sequence was generated and the reads were mapped to the new consensus again. The procedure continued until the consensus converged on a single sequence.
All rabies reads were initially mapped to GenBank rabies reference sequence GI:260063801. This reference sequence was used as the common coordinate system for comparing samples and identifying coding frames. Similarly, GenBank bovine coronavirus GI:15081544 was used as the reference sequence for the BCV samples.
Based on a later observation that our sequenced rabies virus genome differed by approximately 9% relative to the pre-selected reference fox rabies genome, we checked to see if observed error rate (defined below) would increase by introducing random mutations at 9% of the plasmid control reference sequences generated from Sanger sequencing. Increased divergence between the sample and the randomly mutated reference sequence could confound the read mapping program and introduce additional alignment errors, however, no noticeable increase in error rates were observed, suggesting that the read mapping parameters were able to tolerate this rate of divergence.
Estimation of error rate from control plasmids
The two plasmid controls were used to empirically model combined PCR and sequencing errors as well as to evaluate our algorithm for making genetic variant calls. The clone control samples were amplified using the same PCR amplification protocol as was used for sequencing natural samples. A control reference sequence was generated from a separate Sanger sequencing run. Any polymorphisms that deviated from the consensus sequence were taken to be examples of error introduced either through PCR amplification or sequencing.
At every base, any nucleotide called by a read is referred to as a “candidate base call”. Error rates were calculated as a ratio between the total number of candidate base calls differing from the consensus nucleotides summed across the genome and the total number of base calls made across the genome.
Quality control of the sequencing data
The following rules were implemented to maximize the quality of the reads. Quality scores (Q-score) from the sequencer were used to compile nucleotide frequency distribution for every base sequenced. These nucleotide frequency distributions were generated at four quality thresholds for comparative analyses: raw (all reads), Q≥10, Q≥20 and Q≥30. At a given base, a read covering the base contributes to a candidate base call only when the minimum Q-score over an 11-nucleotide window (±5bp) centered on the query base surpasses the quality-score threshold being considered (e.g. the Q30 nucleotide frequency profile). In addition to Q-score filtering, a misalignment filter required that the 11-nt window contained no indels and that the query position must be at least five bases away from the end of the read to avoid misalignment for single reads. To avoid the potential of higher error rates at the 3-prime ends of single reads, only the first 80 bases were used in single read analyses. Furthermore, when calculating error rates and making variant calls, only those bases where greater than 10% of the reads that survive the quality filters are considered for analysis at that quality score. This is to avoid inclusion of non-representative features for select bases in the genome.
Sequencing error analysis
At every base, all overlapping read pairs were separated into two categories: matching and non-matching base pairs. Matching base pairs have two complementary nucleotides and non-matching base pairs have two incongruent nucleotides. Mismatched read pairs were only used to calculate mismatch rates and examine the relationship between mismatch rates and quality scores. Other than that, mismatched read pairs were excluded from all error and variant analyses.
Mismatch rates were calculated two ways, ‘per position’ and ‘per base pair’. Per-position mismatch rate is the fraction of overlapping read pairs that are mismatched at any given location in the genome. Per base pair mismatch rate is the total number of mismatched read pairs summed across all bases in the genome divided by the total number of read pairs.
To generate quality score distributions for the matching and mismatched read pairs, Q-scores for every base pair were compiled as follows. For a matching base pair, the average quality score was used. For a non-matching base pair, the minimum quality score was used. The resulting Q-score distributions were compared and used to generate Q-score receiver-operator characteristic (ROC) and ‘false discovery rate’ curves.
Sequencing errors can occur in two forms in overlapping read pairs. Non-complementarity between the forward and reverse strands at a given base indicates that at least one of the two nucleotides is erroneously incorporated. This type of error is straightforward to exclude -- all non-matching read pairs are excluded from analysis except in the analyses for quality control. A second, more rare but ‘hidden’ form of error is where two complementary errors occur on both the forward and the reverse strands such that the resulting read pair remains complementary.
Making variant calls
The error rate, p, is the combined PCR and sequencing error, ε, adjusted by a function of the ORP mismatch rate, δ, at the base in question.
P-value = 0.01 with Bonferroni correction was used as the significance threshold for each hypothesis test.
Scripts for the variant detection model were written in Python and R and are available upon request.
This work was performed under the auspices of the U.S. Department of Energy by Lawrence Livermore National Laboratory under Contract DE-AC52-07NA27344. This work was funded in part from a grant from the Defense Threat Reduction Agency. We thank Sharon Messenger from California Department of Public Health for providing the rabies samples and Pamela Hullinger for assisting in the collection of BCV samples. We thank Daniel Newburger and Hugo Lam for their critique on the manuscript.
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