DOI: 10.5061/dryad.1vhhmgqtx
title: A quantitative analysis of vertebrate environmental DNA degradation in soil in response to time, UV light and temperature

description: Environmental DNA (eDNA) degradation influences the effectiveness of
 eDNA-based biodiversity monitoring, but the factors that determine the
 rate of decay of eDNA in terrestrial environments are poorly understood.
 We assessed the persistence of vertebrate eDNA from a mock vertebrate
 community created with soil from zoo enclosures holding ten target species
 from different taxonomic classes (e.g. reptiles, birds and mammals) and of
 different biomass (e.g. little penguin and giraffe). We examined species
 detection rates resulting from eDNA metabarcoding, as well as relative
 eDNA concentrations via qPCR, from soil samples over eight time points (0
 to 12 weeks), during exposure to three ambient temperatures (10, 25 and 40
 °C) and three levels of ultraviolet B (UV-B) radiation (0%, 50% and 100%
 intensity). We recorded considerable variation in detectability between
 species, independent of temperature and UV-B effects. Quantitative PCR
 indicated degradation of eDNA over time for all temperature and UV
 treatments, although it was still possible to detect eDNA from some
 species after twelve weeks. Degradation rates were lowest for high UV-B
 treatments, presumably due to UV-B reducing bacterial metabolism. The
 temperatures investigated did not influence eDNA decay. Our results
 indicate that eDNA in soil can persist under a range of temperatures and
 high UV radiation for longer than expected. Sheltered sites with minimal
 UV-B radiation, which have previously been considered ideal sites for
 terrestrial eDNA collection, may not be optimal for eDNA persistence in
 some cases due to microbial decay. A better understanding of eDNA
 degradation in terrestrial environments is needed before eDNA
 metabarcoding of soil can be used as a stand-alone method for accurate
 surveying of terrestrial vertebrate communities.
description: <strong>Sample source</strong>
 Soil was collected in mid-July 2021 from the enclosures of ten
 species at the Perth Zoo, South Perth, Western Australia. Soil samples
 (1.5 L), consisting of five equal subsamples from areas of recent animal
 activity (as determined by zoo staff) to maximise the likelihood of eDNA
 presence in a sample, were collected from each of four mammal, three bird
 and three reptile enclosures (Table 1): approximately 15 L of soil in
 total. Samples were taken from a depth of 0-50 mm, using sterilised
 sampling equipment and disposable nitrile gloves. Soil texture varied from
 sand (e.g. Cape porcupine) to sandy clay loam (e.g. Galápagos
 tortoise). Target species (Table 1) did not occur in
 the Perth metropolitan region, meaning that any DNA detected from species
 in the samples must be derived from the original soil samples, not
 reintroduced through subsequent contamination. The emu (<em>Dromaius
 novaehollandiae</em>) was a possible exception, though contact with
 DNA from this species was deemed to be highly unlikely between sampling
 events. A mock community of zoo vertebrate eDNA was
 created by homogenising the ten soil samples using a combination of soil
 sieves and mixing. Equal aliquots of the resulting soil mix were placed in
 aluminium trays (26.5 x 19.5 x 6.5 cm; approximately 550 mL per tray at 10
 mm thickness) and exposed to three levels of ambient temperature (10, 25
 and 40°C) and three levels of UV light intensity (0%, 50% and 100%), with
 three replicates for each treatment (27 trays in total). Modified plant
 growth cabinets (Arcus Refrigeration) maintained constant temperature and
 UV conditions for the duration of the experiment. Ultraviolet light was
 produced by 26-watt ExoTerra Reptile UVB200 bulbs, producing approximately
 240 µW/cm<sup>2</sup> and 40 µW/cm<sup>2</sup> of
 UV-A and UV-B respectively (ExoTerra, 2021).  Temperature levels were
 selected to represent typical ambient air temperatures occurring in
 terrestrial settings across a variety of Australian climatic zones, after
 Jones et al. 2009. iButton temperature loggers showed that the temperature
 of soil in treatment trays was in equilibrium with climate control chamber
 ambient temperatures. Clear plastic wrap was used to cover the 100% UV
 treatment trays and did not affect UV transmission (measured using UV-vis
 spectrometry - see Supplementary Figure 1). 50% UV treatment trays were
 covered with clear plastic wrap and 50% UV-block shade cloth. Zero UV
 light treatment trays were covered with aluminium foil. All trays were
 well sealed on all edges between sampling points. Soil samples were
 collected in 50 mL falcon tubes from the surface soil (&lt;10 mm) of
 each tray as the experiment commenced (Time 0), with subsequent sampling
 at 6 h, 24 h, 48 h, 1 week, 3 weeks, 6 weeks and 12 weeks later. Each
 sample was taken from a different area of the tray. Samples were stored at
 -21.0 °C until DNA extraction. <strong>Sample
 Processing</strong> <em>DNA
 extraction</em> Soil DNA extraction and PCR were
 conducted at TrEnD (Trace and Environmental DNA) facilities at Curtin
 University, Perth, Western Australia. Samples were homogenised using a
 Qiagen TissueLyser, and DNA extracted from 250 mg of soil using the DNeasy
 PowerLyser Powersoil kit (Qiagen), followed by the Qiacube extraction
 platform (Qiagen). Extracts were kept at -20.0 °C until use. The final
 elution volume was 100 μL, and extraction controls (blanks, n = 10) were
 processed for every set of extractions.
 <strong>PCR amplification</strong>
 Quantitative Polymerase Chain Reaction (qPCR; Applied Biosystems,
 USA) was used to assess the quality and quantity of the DNA extracts and
 determine the optimum DNA input (Murray et al 2015).  The
 vertebrate-specific metabarcoding assay 12S-V5 (Riaz et al., 2011) was
 used, targeting a 98-bp region of the mitochondrial 12S gene with the
 primers F2 (5’-TAGAACAGGCTCCTCTAG-3’) and R2 (5’-TTAGATACCCCACTATGC-3’).
 Extracts were amplified on a StepOnePlus Real-Time PCR System (Applied
 Biosystems) under the following conditions: 95 °C for 5 minutes, and 50
 cycles of 95 °C for 30 seconds, 58 °C annealing temperature for 30 seconds
 and 72 °C for 45 seconds. A melt curve of 95 °C for 15 seconds, 60 °C for
 one minute and 95 °C for 15 seconds was used, ending with ten-minute
 elongation at 72 °C. Each 25 µL qPCR mix contained 2.5mM
 MgCl<sub>2</sub> (Applied Biosystems), 1× PCR Gold buffer
 (Applied Biosystems), 0.25 mM dNTPs (Astral Scientific, Australia), 0.4
 mg/mL bovine serum albumin (Fisher Biotec, Australia), 0.4 μmol/L forward
 and reverse primer, 1 U AmpliTaq Gold DNA polymerase (Applied Biosystems)
 and 0.6 μL of a 1:10,000 solution of SYBR Green dye (Life Technologies,
 USA). Cycle-threshold (C<sub>t</sub>) values were recorded for
 all samples and dilutions. PCR reactions were performed with two dilutions
 (3 PCR reactions per extract at neat, 1:10 and 1:100), and the optimum
 level of DNA input was used for fusion tag PCR. Extraction controls,
 positive controls (<em>Gallus gallus</em>) and non-template
 controls were included in qPCR runs. All PCR mixes were
 prepared in a dedicated laboratory to minimise contamination, and DNA
 extracts were added in a separate laboratory in dedicated UV cabinets.
 Samples containing amplifiable DNA as determined by qPCR were assigned a
 unique combination of fusion tag primers, containing unique multiplex
 identifier (MID) tags with lengths between 6 and 8 bp, the 12S-V5 primer
 and sequencing adapters (Illumina). A single step fusion protocol was used
 with no reuse of index combinations, and samples were amplified in
 duplicate using the same conditions as the qPCR protocol described
 previously, reducing the effect of PCR stochasticity (Murray et al.,
 2015). Duplicate samples were pooled alongside amplicons with similar ΔRn
 values. Pooled amplicons were cleaned using the QIAquick PCR purification
 kit (Qiagen) and quantified using the QIAxcel Advanced system
 (Qiagen). <strong>DNA library preparation and
 sequencing</strong> A DNA library was created by
 combining pools in equimolar ratios based on the quantification, and then
 size selected (160-400 bp) using Pippin Prep (Sage Sciences) to remove
 MID-tag primer-dimer products formed in amplification. The final library
 was cleaned using a QIAquick PCR purification kit (Qiagen) and eluted to
 50 µL. A QuBit fluorometer (Thermo Fisher Scientific) was used to quantify
 the final DNA library, before sequencing on Illumina’s MiSeq platform
 (Illumina, San Diego, CA, USA) using a single-lane flow cell as per
 Illumina protocols for single-end sequencing (MiSeq v2 Reagent Kit 300
 Cycles). <strong>Sequence
 analysis</strong> Raw sequence reads were
 processed using the eDNAFlow pipeline (Mousavi-Derazmahalleh et al.,
 2021). Sequences were quality-filtered using AdaptorRemoval v2 (Schubert
 et al., 2016) and FASTQC (Andrews, 2010), and demultiplexed using OBITools
 (Boyer et al., 2016). Reads were dereplicated, denoised and assigned a
 zero-radius operational taxonomic unit (ZOTU) using Usearch (Edgar, 2010),
 with sequences shorter than 60 bp removed. Taxonomic assignment was
 conducted by matching ZOTU sequences to Genbank, an online reference
 database (https://www.ncbi.nlm.nih.gov/genbank/), using a basic local
 alignment search tool (BLASTn) run on a high-throughput HPC cluster
 (Pawsey Supercomputing Centre Perth, WA, Australia). Final taxonomic
 assignments were conducted using a custom lowest common ancestor (LCA)
 algorithm in Python (Mousavi-Derazmahalleh et al., 2021), requiring 95%
 query coverage and 90% identity match with reference sequences. Where the
 LCA script was unable to assign a ZOTU to a species level or the species
 detected was not found in the study area, genus level was reported, and
 the ZOTU assessed manually based on known occupants of both the Perth Zoo
 and surrounding area. Samples with a low sequencing depth (less than 5000
 total reads, as determined by rarefaction curves; Supplementary Figure 2)
 were removed from the data, and individual ZOTU detections with less than
 two sequence reads were removed. Due to the minimal contamination risk
 associated with single-fusion protocols through physical separation of pre
 and post-PCR areas, good laboratory practices and no reuse of fusion tag
 combinations (Taberlet et al., 2018), any detections above two identical
 sequence reads were considered ‘real’ detections. ZOTUs detected in
 positive or negative controls, as well as those from common PCR
 contaminants (e.g., human, ungulate species) were removed prior to
 analysis. C<sub>t</sub> values from qPCR runs were recorded
 for each undiluted DNA extract and compared for sampling time and
 temperature and UV exposure. These values corresponded to the relative
 concentrations of vertebrate eDNA within each extract. Lower
 C<sub>t</sub> values (less cycles to amplify DNA over a
 threshold) represented higher relative DNA concentrations, whilst higher
 C<sub>t</sub> values signified lower relative DNA
 concentrations.
description: # A quantitative analysis of vertebrate environmental DNA degradation in
 soil in response to time, UV light and temperature
 [https://doi.org/10.5061/dryad.1vhhmgqtx](https://doi.org/10.5061/dryad.1vhhmgqtx) ## Description of the data and file structure There are six files included: **MSRun480-FTP360_S1_L001_R1_001.fastq.gz** - Compressed sequencing file containing raw sequences from the Illumina MiSeq run **MSRun485-FTP365_S1_L001_R1_001.fastq.gz** - Compressed sequencing file containing raw sequences from the Illumina MiSeq run **MSRun504-FTP375_S1_L001_R1_001.zip** - Compressed sequencing file containing raw sequences from the Illumina MiSeq run **Sample_barcode_eDNA_degradation.csv** - sample metadata including information required to demultiplex raw sequences **Sample_CT_values.csv** - CT values recorded for soil samples **filtered_ZOTU_table_Zoo.csv** - Final ZOTU table produced using eDNAFLow bioinformatics pipeline for sequence analysis. Taxonomy included in this file and bioinformatics steps performed are further described in the methods section of the publication. Details for **Sample_barcode_eDNA_degradation.csv** MiSeq_Run - ID of the MiSeq Run MiSeq_Run_Date - Date of the sequencing run Sample.ID - Sample ID Description - Type of sample taken Site - Sample site Extraction - Extraction kit used F_Tag_ID - ID of the Forward multiplex identifier (MID) tag R_Tag_ID - ID of the Reverse multiplex identifier (MID) tag F_Primer, R_Primer - IDs of the forward and reverse primers F_Tag_seq - sequence of the Forward multiplex identifier (MID) tag R_Tag_seq - sequence of the Reverse multiplex identifier (MID) tag F_Primer_Seq - Forward Primer sequence R_Primer_Seq - Reverse Primer sequence Primer - Name of the assay used Details for **Sample_CT_values.csv** ID - Soil tray ID with 'TEMPERATURE_UV_REPLICATES'.\ Temperature - ambient temperature of the plant growth chamber in degrees Celcius.\ UV - UV intensity of each foil tray (L = low, M = medium, H = high).\ Replicates - biological replicate number for each set of conditions.\ columns T0-T12W - CT (cycle threshold) values for each sample from time 0 to time 12 weeks. Details for **filtered_ZOTU_table_Zoo.csv** First 7 columns (domain, phylum, class, order, family, genus, species) correspond to the classification of each ZOTU.\ taxon - simplified classification to lowest taxonomic resolution.\ OTU - Operational Taxonomic Unit, a clustered set of DNA sequences obtained from environmental samples, representing a single, distinct taxon.\ All other columns are samples with unique sample IDs.