Data for: Validation of a nutria (Myocastor coypus) environmental DNA assay highlights considerations for sampling methodology
Data files
Mar 29, 2023 version files 37.35 KB
Abstract
Nutria (Myocastor coypus) is a semi-aquatic rodent species that is invasive across multiple regions within the United States. Here we evaluated a qPCR assay previously described for use in Japan for application across invasive populations in the United States. We also compared two environmental DNA sampling methodologies for this assay: field filtration of large volumes of water passed through filters versus direct sampling of small volumes of water. We validated assay specificity, generality, and sensitivity, compared assay performance between two independent laboratories, and successfully tested the assay in situ on a known wild population. The filtration method required fewer samples for environmental DNA detection than direct sampling, but the choice of methods should be assessed based on specific field conditions and time and budget considerations. Our extensive assay validation and comparison across laboratories suggests that the assay is ready to be applied in environmental DNA monitoring of nutria throughout the United States.
Methods
We evaluated assay specificity in vitro by testing DNA extracted from tissue, blood, or scat samples from one individual of each of six related nontarget sequences, including guinea pig, long-tailed chinchilla, North American porcupine , muskrat, mountain beaver, and North American beaver. Tissue samples were sourced from collections at the National Genomics Center for Wildlife and Fish Conservation (NGC; Missoula, MT). These samples were originally collected by state and federal partners from wildlife mortalities in accordance with appropriate sampling permits and wildlife handling approvals. Scat samples from guinea pig and long-tailed chinchilla were collected from animal pens at a local pet store and did not require any animal handling. DNA from all samples was extracted via DNeasy Blood and Tissue Kits (QIAGEN) and diluted to 0.1 ng/µL, as determined using a Qubit 2.0 fluorometer (Thermo Fisher Scientific) prior to quantitative polymerase chain reaction (qPCR). Each qPCR reaction was 15 µL total, composed of 7.5 μL of 2× TaqMan Environmental Mastermix 2.0 (Life Technologies), 0.75 μL of 20x assay mix (600 nM of each primer and 250 nM of probe), 4 μL template DNA, and 2.75 μL of laboratory‐grade sterile water. Primer concentrations in the above reaction were chosen by testing all 16 possible combinations of forward and reverse primers at a concentration of 100, 300, 600, and 900 nM; the optimal concentration combination (600:600) was the concentration producing the lowest quantification cycle (Cq) value and highest end-point fluorescence with a target synthetic gene. Thermocycling conditions were 95°C 10 min, [95°C 15 s, 60°C 60 s] × 45 cycles, as outlined by Akamatsu et al. (2018). We multiplexed the assay with an internal positive control (TaqMan Exogenous Internal Positive Control; Life Technologies) to test for the presence of PCR inhibitors. On each qPCR plate, we included a negative control well and nutria tissue extracts as positive control wells to ensure that the assay was performing as expected. We considered a positive as linear amplification, determined by expert technician judgement, prior to 45 cycles with typical curve morphology. We considered a positive in any qPCR well as a detection.
We evaluated assay generality in vitro by testing genomic DNA from 50 nutria collected throughout four distinct regions of their invasive range in the United States: 15, 5, 20, 8, and 2 from California, Louisiana, Maryland, Oregon, and Virginia, respectively. Additionally, a representative tissue sample from each state was Sanger sequenced in-house or via Eurofins Genomics (Louisville, Kentucky, USA; Supporting Informaiton Appendix A). Sequences were aligned and edited using Sequencher v5.4.6 (Gene Codes, USA) and compared to the nucleotide database on GenBank using default settings in Basic Local Alignment Search Tool (BLAST; February 2023). Tissues were collected from euthanized nutria that were sampled ancillary to population reduction for damage mitigation conducted by the United States Department of Agriculture (USDA) Animal and Plant Health Inspection Service (APHIS) Wildlife Services and the California Department of Fish and Wildlife. Given that tissue samples were collected secondarily to legally authorized control of nutria, sample collection was exempted from Institutional Animal Care and Use Committee review. Tissues were stored, processed, and analyzed as described above.
We compared limit of detection (LOD) and limit of quantification (LOQ) across two laboratories—the NGC and the National Wildlife Research Center (NWRC; Fort Collins, CO)—to evaluate the consistency of assay performance. Both laboratories determined LOD and LOQ via qPCR with replicate standard curves derived from synthetic double-stranded DNA of nutria (gBlock gene fragments; Integrated DNA Technologies). Specifically, the NGC laboratory analyzed LOD and LOQ using a single plate with 6 replicates of each of 7 dilution levels (2, 10, 50, 250, 1,250, 6,250, and 31,250 copies per reaction) and 6 no template controls. The NWRC laboratory analyzed 3 plates, each with 16 replicates of each of 5 dilution levels (n = 48 each at 0.4, 2, 4, 40, and 400 copies per reaction), 8 positive controls, and 8 no template controls (n = 24 each). We analyzed results using the curve fitting methods described by Klymus et al. (2019) to determine LOD and LOQ. We used the option “Best” in the corresponding R code to analyze multi-lab LOD/LOQ study data to automatically select the best fitting model choice for LOD and the model with the lowest residual standard error for LOQ (R Core Team, 2021; Klymus et al. 2019; Merkes et al. 2019).
We tested the assay in situ using environmental (i.e., water) samples collected in Oregon, USA, from areas with known presence of nutria (n = 9 waterbodies; nutria observed or fresh sign reported by the collector; collected from Marion County in November 2021). We compared two DNA collection and extraction methods using known-present samples. The first method, hereafter referred to as the filtration method, followed the protocol of Carim, McKelvey, Young, Wilcox, and Schwartz (2016). To isolate DNA from the filters, we cut each filter in half; one half of the filter was archived at -20 °C for future analyses and the other half was extracted using a modified DNeasy Blood and Tissue Kit (QIAGEN) protocol. We followed the manufacturer’s protocol for extraction from animal tissue with the modifications described in Carim, Dysthe, Young, McKelvey, and Schwartz (2016). In cases where a sample consisted of multiple filters, we extracted each filter separately and then spun them through the same silica spin column (see Carim, Dysthe, et al. 2016). We stored DNA extractions at -20 °C until analysis. Samples from areas with known absence of nutria (n = 7; site elevation is outside the range of nutria cold tolerance; collected in Union County in October 2017) were also collected and extracted using this filtration method. The second DNA collection and extraction method, hereafter referred to as the direct sampling method, followed the optimized protocol of Williams, Huyvaert, & Piaggio (2017). We collected three water samples from each site and two negative controls (i.e., bottled water with Longmire’s buffer) to evaluate potential contamination. We stored water samples at -20 °C prior to receipt at the NWRC laboratory where they were stored at -80 °C prior to analysis. For DNA capture, two subsamples (15 mL each) were centrifuged at 9,000 g for 15 minutes and DNA was isolated from each pellet with the DNeasy mericon Food Kit (QIAGEN) following the manufacturer’s Standard Protocol (200 mg). DNA extracts and the remaining 30 mL of each water sample were stored at -80°C. To analyze environmental samples, we used the qPCR condictions described in Akamatsu et al. (2018) and outlined above with the addition of an internal positive control to test for the presence of PCR inhibitors, defined as an internal positve control in the sampling being shifted > 1 Cq higher than that in the negative control. We multiplexed the assay with an internal positive control (TaqMan Exogenous Internal Positive Control; Life Technologies) by replacing 1.8 μL of water with 1.5 μL of 10X exogenous IPC assay and 0.3 μL of 50X exogenous IPC DNA per reaction. We analyzed each extraction with three qPCR replicates and on each qPCR plate we included three negative control wells and a dilution series of synthetic gene fragments as positive controls.
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