DOI: 10.5061/dryad.jq2bvq8fh
title: An examination of seasonal variation in taxonomic richness and community composition using eDNA on a tropical coral reef

description: Small volumes of water containing environmental DNA (eDNA) are
 increasingly combined with metabarcoding to generate biodiversity data for
 specific fractions of marine flora and fauna. To date, however, few
 studies have utilized this technique to assess how well it captures
 seasonal patterns in coral reef communities or how environmental or
 methodological factors influence eDNA detections. In our study, we used
 three eDNA metabarcoding assays primarily targeting bony fish,
 elasmobranchs, as well as cnidarians and sponges (Cnidaria/Porifera)
 combined with monthly seawater sampling to 1) investigate temporal
 variation in taxonomic detections and 2) statistically test the potential
 effect of season, sea surface temperature, timing of spawning (using moon
 phase as a proxy), and sample preservation on taxon detection across a
 12-month period in a model coral reef system (Big Vicki’s Reef, Lizard
 Island, Great Barrier Reef, Australia). Species-level fish and genus-level
 scleractinian coral detections from standardised visual surveys conducted
 at the same coral reef, in addition to a curated list of all known fishes
 recorded from the more expansive coral reef system, were used to validate
 eDNA detections. Our eDNA dataset indicated that the number of taxa
 detected were consistently highest in September for fish, and in February
 followed by September for Cnidaria/Porifera. Conversely, detections were
 lowest in June and July for all taxa. Some, but not all, of the
 environmental and methodological variables explained the observed temporal
 pattern in biological communities or systematic changes in the number of
 taxa, and in some cases, this effect was taxon dependent. Our study also
 highlights the significance of timing in eDNA biodiversity surveys
 conducted on tropical coral reefs in the Southern Hemisphere. To obtain
 the most meaningful estimates of site diversity, we recommend focusing
 sampling efforts between early spring to early autumn. Alternatively,
 allocating an entire year to sampling would better capture seasonal
 variation and provide more comprehensive insights into coral reef
 biodiversity.
description: <strong>Laboratory processing</strong>
 DNA was extracted from half of each filter membrane (including
 filtration controls) within six months of collection using the DNeasy
 Blood and Tissue Kit (QIAGEN) in the TrEnD Laboratory at Curtin University
 in Western Australia with the following modifications: 540 µl of ATL lysis
 buffer, 60 µl of Proteinase K, and a 3-hr digest at 56°C. An extraction
 blank was processed in parallel with every set of eleven samples to detect
 any cross-contamination. Three previously published primer sets (16S Fish,
 CoralITS2, and CoralITS2_acro; Table 1) were employed in this study to
 amplify primarily teleost fish and scleractinian coral taxa, respectively.
 Quantitative PCR (qPCR) was carried out in 25 μl containing the following
 concentrations: 1X AmpliTaq Gold® PCR buffer (Life Technologies,
 Massachusetts, USA), 2 mM MgCl2 (Fisher Biotec, Australia), 0.4μM dNTPs,
 0.1mg BSA (Fisher Biotec, Australia), 0.4 μM each of forward and reverse
 primers (Integrated DNA Technologies, Australia), 0.6 μl of 5X SYBR® Green
 (Life Technologies), 1U AmpliTaq Gold® DNA Polymerase (Life Technologies),
 2 μl of eDNA template, and made to volume with Ultrapure™ Distilled Water
 (Life Technologies). All qPCRs were prepared in dedicated trace DNA (clean
 room) facilities at the TrEnD Laboratory, Curtin University, and amplified
 on a StepOnePlus Real-Time PCR System (Applied Biosystems, Massachusetts,
 USA) with the following PCR cycling conditions: initial denaturation at
 95°C for 5 min, followed by 40 cycles of 95°C for 30 s, 52-55°C for 30 s
 (see respective annealing temperatures in Table 1), and 72°C for 45 s,
 with a final extension of 72°C for 10 min. Each sample was amplified in
 duplicate in a single-step process via the use of fusion-tagged primer
 architecture, whereby original primers are flanked by a unique 6–8 bp
 multiplex identifier tag (MID-tag) and an Illumina compatible sequencing
 adapter.  MID-tagged PCR amplicons were then pooled
 into two libraries (the first containing 16S Fish amplicons, and the
 second CoralITS2 and CoralITS2_acro amplicons combined) at equimolar
 ratios based on qPCR ΔRn values. The two libraries were then size-selected
 using a Pippin-Prep (Sage Science, Beverly, USA) to remove any amplicons
 outside of the target range (150-450 bp, and 160-600 bp, respectively).
 Size-selected libraries were then purified using the QIAquick PCR
 Purification Kit (Qiagen, Venlo, Netherlands), quantified using a Qubit
 4.0 Fluorometer (Invitrogen, Carlsbad, USA), and diluted to 2 nM for
 loading onto the sequencing platform. The 16S Fish library was loaded onto
 a 300 cycle Illumina MiSeq® V2 Standard Flow Cell for unidirectional
 sequencing. The CoralITS2 library was loaded onto a 500 cycle Illumina
 MiSeq® V2 Standard Flow Cells for paired-end sequencing. The sequencing of
 both libraries was conducted on an Illumina MiSeq platform (Illumina, San
 Diego, USA) housed in the TrEnD Laboratory.
 <strong>Bioinformatics</strong>
 Sequences were demultiplexed into their respective samples based
 on their MID-tags using the <em>ngsfilter</em> and
 <em>obisplit</em> commands in OBITools (v1.2.9; Boyer et al.
 2016) for the single-end (16S Fish) sequencing reads, and the
 <em>insect</em> package (Wilkinson et al. 2018) for the
 paired-end (CoralITS2 and CoralITS2_acro) sequencing reads. Data from the
 two sequencing libraries were separately quality filtered (minimum
 sequence length = 100 bp, maximum expected errors = 2, no ambiguous
 nucleotides), denoised, merged (paired-end reads only, 20 bp overlap, no
 mismatches), filtered for chimeras and dereplicated (pool = TRUE) using
 the <em>DADA2</em> package (Callahan et al. 2016) in R
 (v3.5.3; R Core Team 2015). To maximize biodiversity detections, we
 retained high-quality singletons (that passed strict quality filtering,
 denoising, and chimera removal) if they were observed in at least two
 independent samples (DADA2 Pool=TRUE function). If the singleton was only
 found in one sample, it was removed from the dataset. This generated an
 amplicon sequence variant (ASV) .fasta file and count table; the former
 was queried against NCBI’s GenBank nucleotide database (accessed September
 2020; Benson et al. 2005) using BLASTn (minimum percentage identity = 90,
 maximum target sequences = 10, reward value = 1) with Zeus, an SGI
 cluster, based at the Pawsey Supercomputing Centre in Kensington, Western
 Australia. Taxonomic assignments of ASVs were curated and, when necessary,
 collapsed to the lowest common ancestor (LCA) using a custom LCA python
 script (Mousavi‐Derazmahalleh et al. 2021), with a threshold query
 coverage (qCov) of 100%, a minimum percentage identity of 90%, and a
 difference (Diff) of 1. ASVs detected in extraction blanks with a read
 count &gt;10 were removed across the entire dataset, prior to ASVs
 being merged by taxonomy using the phyloseq v1.24.2
 ‘<em>tax_glom</em>’ function (McMurdie and Holmes 2013) in R.
 Sample replicates were then pooled by their temporal sampling period and
 taxa accumulation curves were plotted by sequencing depth using the vegan
 v2.5-7 <em>rarecurve</em> function (Oksanen et al. 2019) in R.
 These rarefaction analyses (Fig. S2 and S3 in Online Resource 1) indicated
 that subsampling of sequencing depth was not required, given that a
 plateau was apparent in the taxa accumulation curves.
description: We here provide the demultiplexed 16S Fish (read 1 only), as well
 as the Coral ITS2 (CP1) and Coral ITS2_acros (CP2; read 1 and read 2)
 assay sequence data in .fastq format, along with an excel spreadsheet
 listing the metadata associated with each seawater sample collected at
 Lizard Island, Australia. These sequences and metadata were used in the
 following paper: DiBattista, J.D., West, K.M.,
 Ceccarelli, D.M., Hoggett, A.K., Vail, L.L., Garcia, R., and Richards,
 Z.T. (in press)<strong> </strong>An examination of seasonal
 variation in taxonomic richness and community composition using eDNA on a
 tropical coral reef. <em>Coral Reefs</em>. https://doi.org/10.1007/s00338-024-02594-6
description: # An examination of seasonal variation in taxonomic richness and community
 composition using eDNA on a tropical coral reef \[Access this dataset on
 Dryad
 (doi:[https://doi.org/10.5061/dryad.jq2bvq8fh](https://doi.org/10.5061/dryad.jq2bvq8fh))] ## Description of the data and file structure We here provide the raw demultiplexed 16S Fish (read 1 only), as well as the paired-end Coral ITS2 (CP1) and Coral ITS2_acros (CP2) assay sequence data in .fastq format. These files contain sequence data seperated into their individual sample replicates for each genetic assay. For file naming nomenclature, each file name begins with the acronym for Lizard Island Research Station (LIRS), followed by the date of sampling, followed by the replicate number, followed by the genetic assay, and in the case of paired-end sequencing runs/assays (i.e. CP1 and CP2), whether it is the read 1 set of sequences (R1) or the read 2 set of sequences (R2). For example, LIRS_12_01_20_1_16S_Fish.fastq are the DNA sequences in .fastq format obtained from the water sample collected at LIRS, on January 12, 20220, replicate 1 (of 6) samples collected on that date, and sequenced using the 16S Fish genetic assay. As another example, LIRS_12_02_19_3_CP1_R1.fastq are the DNA sequences in .fastq format obtained from the water sample collected at LIRS, on February 12, 2019, replicate 3 (of 6) samples collected on that date, run on the Coral ITS2 (CP1) genetic assay, and the R1 set of sequences. As a final example, LIRS_12_02_19_3_CP1_R2.fastq are the DNA sequences in .fastq format obtained from the water sample collected at LIRS, on February 12, 2019, replicate 3 (of 6) samples collected on that date, run on the Coral ITS2 (CP1) genetic assay, and the paired R2 et of sequences. We also provide an excel spreadsheet listing the metadata associated with each seawater sample collected at Lizard Island, Australia named "LI_sampling_metadata__Final__July_7__2023.csv" EMu registration number: Australian Museum voucher registration number for the archived filter paper. Sample ID: unique sample ID based on the acronym for Lizard Island Research Station (LIRS), followed by the date of sampling, followed by the replicate (filter paper) number. Amount filtered: amount of water pushed through each replicate filter paper in litres. Filter size: mesh size of the filter paper used to filter water samples. Sample processing: the person that filtered the water sample. Sampling country: country where water sample was collected. Sampling Location: exact location where the water sample was collected. Reef Location: broader reef where the water sample was collected. Exposed or Sheltered: whether water sample was collected on the exposed or sheltered side of the reef. Latitude: latitude where the water sample was collected. Longitude: longitude where the water sample was collected. Sampling Date: date when the water sample was collected Lunar Cycle: lunar position when the water sample was collected (a variable used to test associations with taxonomic richness and composition) Days frozen: number of days the collected water sample was frozen prior to thawing and filtering. Note that 0 days frozen means the water sample was not frozen but instead filtered immediately after collection on the reef. ## Sharing/Access information These sequences and metadata were used in the following published paper: DiBattista, J.D., West, K.M., Ceccarelli, D.M., Hoggett, A.K., Vail, L.L., Garcia, R., and Richards, Z.T. (in press) An examination of seasonal variation in taxonomic richness and community composition using eDNA on a tropical coral reef. Coral Reefs. [https://doi.org/10.1007/s00338-024-02594-6](https://doi.org/10.1007/s00338-024-02594-6)