DOI: 10.5061/dryad.vq83bk3wr
title: Data for: Validation of a nutria (Myocastor coypus) environmental DNA assay highlights considerations for sampling methodology

description: 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.
description: 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 &gt; 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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