DOI: 10.5061/dryad.8cz8w9gng
title: Money spider dietary choice in pre- and post-harvest cereal crops using metabarcoding

description: Money spiders (Linyphiidae) are an important component of conservation
 biological control in cereal crops, but they rely on alternative prey when
 pests are not abundant, such as between cropping cycles. To optimally
 benefit from these generalist predators, prey choice dynamics must first
 be understood. Money spiders and their locally available prey were
 collected from cereal crops two weeks pre- and post-harvest. Spider gut
 DNA was amplified with two novel metabarcoding primer pairs designed for
 spider dietary analysis, and sequenced. The combined general and
 spider-exclusion primers successfully identified prey from 15 families in
 the guts of the 46 linyphiid spiders screened, whilst avoiding
 amplification of Erigone spp. The primers show promise for application to
 the diets of other spider families such as Agelenidae and Pholcidae.
 Distinct invertebrate communities were identified pre- and post-harvest,
 and changes in spider diet and, to a lesser extent, prey choice reflected
 this. Spiders were found to consume one another more than expected,
 indicating their propensity toward intraguild predation, but also consumed
 common pest families. Changes in spider prey choice may redress prey
 community changes to maintain a consistent dietary intake. Consistent
 provision of alternative prey via permanent refugia should be considered
 to sustain effective conservation biocontrol.
description: Primer development and testing Existing PCR
 primers were tested and ultimately redesigned to better match the target
 taxa of this study. Two novel primer pairs were used for amplification of
 DNA for the dietary analysis of spider gut contents to overcome the
 problems associated with the taxonomic proximity of spiders and their prey
 (particularly other spider species). Novel PCR primers were adapted for
 the exclusion of all spider DNA, with a focus on linyphiids (henceforth
 spider exclusion primers, titled TelperionF-LaurelinR), based upon a
 primer site slightly 3’ of the general animal barcoding primers LCO1490
 (forward primer Folmer <i>et al.</i> 1994), and mICOIintR
 (Leray <i>et al.</i> 2013). A second primer pair was employed
 for broad amplification of both spiders and their prey (henceforth general
 primers, titled BerenF-LuthienR), based upon mICOIintF (Leray <i>et
 al.</i> 2013) and HCO2198 (Folmer <i>et al.</i> 1994).
 Both primer pairs were adapted via base changes designed with reference to
 mass alignments of invertebrate COI sequences and tested <i>in
 silico</i> and <i>in vitro</i>. The spider-exclusion
 primers were designed to overcome the loss of reads to predator DNA,
 whilst the general primers were designed to avoid the taxonomic biases
 associated with the exclusion primers. Mass-alignments
 of COI sequences were batch-downloaded from GenBank (NCBI) and BOLD
 (Ratnasingham and Hebert 2007) using PrimerMiner (Elbrecht and Leese 2016)
 in R v.3.3.4 (R Core Team 2020) to aid visual inspection of existing and
 novel primer sites. PrimerMiner clusters batch downloads into operational
 taxonomic units (OTUs) based on sequence similarity and visualises mass
 alignments of sequence data for primer design. By merging overrepresented
 and duplicate sequences through taxonomy-independent clustering,
 PrimerMiner accounts for within-species variation and cryptic species
 whilst ignoring rare haplotypes (Elbrecht and Leese 2016). Sequences were
 downloaded for all terrestrial invertebrate orders available, and
 consensus sequences were created by clustering these into OTUs for each
 order. The COI sequences were trimmed to include only the Folmer region
 (Folmer <i>et al.</i> 1994) using Geneious R10 (Kearse
 <i>et al.</i> 2012) for subsequent use in PrimerMiner.
 Alignments of prey sequences created via PrimerMiner included cereal crop
 spiders, in order to find primer sites conserved between a wide range of
 potential prey, but different for spiders. Where these sites were 100-400
 base pairs apart on sequences from one another or from existing primer
 sites, they were paired, and primers designed (Table 1, Figures S2-S3).
 Existing general invertebrate primer sites were compared against the
 PrimerMiner alignments to identify any potential improvements to the
 primers for the amplification of cereal spider prey. The coverage of
 primers (% amplified) was determined via PrimerMiner using the same mass
 alignments used for primer design. PrimerMiner uses a taxonomy-independent
 database and accounts for adjacent base mismatches and the position of
 each base in the primer. Primers were also analysed using the online
 ThermoFisher Scientific Multiple Primer Analyzer tool. After the primers
 were deemed successful <i>in silico</i>, they were tested
 <i>in vitro</i>. The primer pairs were
 tested <i>in vitro</i> against a wide range of extracted
 invertebrate DNA including spiders, common spider prey and additional
 invertebrates. For this, invertebrate samples included those collected
 from the field site at Burdons Farm, Wenvoe, South Wales, the study site
 used for subsequent ecological analysis. Invertebrates were found via
 manual searching, collected via aspirator and placed in microcentrifuge
 tubes of 100% ethanol. These were identified at 20-50X magnification using
 a light stereomicroscope and taxonomic keys (Goulet and Huber 1993;
 Roberts 1993; Unwin 2001; Ball 2008; Barber 2008; Duff 2012; Dallimore and
 Shaw 2013). Additional invertebrates and DNA were taken from existing
 archived collections within Cardiff University. Extraction of DNA used
 DNeasy Blood & Tissue Kits (QIAGEN Inc., Chatsworth, CA, USA)
 following the manufacturer’s protocol for animal tissue. For predatory
 invertebrates, DNA was extracted from the lower legs, excluding the femur,
 to avoid the inclusion of prey DNA in the gut diverticulae and leg coxae
 (Macıas-Hernández <i>et al.</i> 2018). To verify successful
 extraction, the DNA and negative controls were amplified via PCR with the
 Qiagen PCR Multiplex Kit (Qiagen) with 95 °C for 15 minutes to activate
 the HotStarTaq<sup>®</sup> DNA polymerase, 35 cycles of 95 °C
 for 30 seconds, 40 °C for 90 seconds and 72 °C for 90 seconds,
 respectively, followed by a final extension at 72 °C for 10 minutes using
 universal invertebrate primers LCO1490 and HCO2198 (Folmer <i>et
 al.</i> 1994). PCR reactions comprised 25 µl reaction volumes
 containing 12.5 µl Qiagen PCR Multiplex kit, 0.2 µmol (0.5 µl of 10 µM
 stock) of each primer, 6.5 µl DNase-free water and 5 µl template DNA.
 Amplification was confirmed by gel electrophoresis.
 Primers were initially tested against a small selection of spider
 and non-spider DNA. Temperature-gradient PCRs were used to determine the
 optimal annealing temperatures for the primer pairs selected, with
 temperatures between 40 °C and 60 °C considered and initial tests starting
 5 °C lower than the mean melting temperature of both primers. Inclusion of
 the Q reagent supplied with Multiplex Kits was trialled for each pair to
 ascertain whether this could improve performance, but was ultimately
 excluded in all cases. PCR conditions were: 95 °C for 15 minutes to
 activate the HotStarTaq<sup>®</sup> DNA polymerase, 35 cycles
 of 95 °C, the annealing temperature and 72 °C for 30, 90 and 90 seconds,
 respectively, and a final extension at 72 °C for 10 minutes. PCR reactions
 comprised 25 µl reaction volumes containing 12.5 µl Qiagen PCR Multiplex
 kit, 0.2 µmol (0.5 µl of 10 µM stock) of each primer, 6.5 µl DNase-free
 water and 5 µl template DNA.  Successful amplification was confirmed by
 gel electrophoresis. Once optimised to amplify a range of non-spider
 species whilst amplifying few spiders, or amplify a broad range of all
 species included, primers were further tested on a broader range of DNA
 (Table S3). The TelperionF–LaurelinR primer pair has
 well-conserved sites, facilitating broad coverage with few degenerate
 bases necessary. The terminal base at the 3’ end of Laurelin, being a
 thymine base, critically mismatches with the guanine base present for most
 spider taxa tested; this should theoretically prevent or at least severely
 reduce amplification of spiders with little cost to amplification breadth
 otherwise. The BerenF–LuthienR pair similarly makes use of conserved
 primer sites employed in other studies but adapted for universal
 amplification of the focal taxa of this study. Field
 collection and identification Linyphiids were visually
 located on transects through two adjacent spring barley fields at Burdons
 Farm, Wenvoe in South Wales (51°26'24.8"N,
 3°16'17.9"W), and collected from occupied webs and the ground,
 in August and September 2017. Each
 transect comprised 4 m<sup>2</sup> searching areas at least 10
 m apart and all available linyphiids were collected. Spiders
 were taken from 20 locations along the aforementioned transects, 10
 pre-harvest and 10 post-harvest. Spiders were collected two weeks prior
 (7-13<sup>th</sup> August) to harvest
 (~20<sup>th</sup> August) of the crop and two weeks after
 harvest (4-8<sup>th</sup> September) in crop stubble and
 placed in 100% ethanol using an aspirator. Ground-active linyphiid spiders
 were collected when webs were not abundant. Spiders were taken to Cardiff
 University, transferred to fresh 100% ethanol, adults identified to
 species-level and juveniles to genus, and stored at -20 °C until
 subsequent DNA extraction. Invertebrate prey
 communities were collected using a converted McCulloch GBV 325 G-vac leaf
 blower suction sampler for 1 minute over 4 m<sup>2</sup> areas
 near to those from which spiders for DNA analysis were collected. Samples
 were taken in transects, with 10 samples each pre- and post-harvest (20
 total), split evenly between two adjacent fields. Invertebrate prey
 community samples were taken approximately 10 m apart, in sites near to
 those from which spiders were collected, with different sites used pre-
 and post-harvest. Invertebrates were killed with ethyl acetate and stored
 in 70% ethanol at -20 °C, as these samples were for measurement of the
 invertebrate community and not for molecular analysis. All invertebrates
 were identified to family level under an Olympus SZX7 stereomicroscope
 using morphological keys, except for springtails of Sminthuroidea
 (Sminthuridae and Bourletiellidae, which were often indistinguishable
 following vacuum sampling and preservation due to the fine features
 necessary to distinguish them) which were left at super-family and mites
 (many of which were immature or in poor condition), which were identified
 to order level. <i>Erigone</i>
 <i>atra </i>and <i>E. dentipalpis</i>
 (Erigoninae), and <i>Tenuiphantes</i>
 <i>tenuis</i> (Blackwall, 1852; Linyphiinae) were the focus of
 our study, although a few juveniles were included from other genera due to
 the difficulties associated with morphological identification of linyphiid
 juveniles; these misidentifications were confirmed in the subsequent
 metabarcoding. In total, 66 spiders were screened (Table S2), unevenly
 split across the 20 corresponding prey sampling sites. Spiders were washed
 in and transferred to fresh 100 % ethanol to reduce external contaminants
 prior to identification using Roberts (1993) morphological key. Abdomens
 were removed from spiders and washed again in fresh 100 % ethanol. Only
 abdomens were used for molecular analysis of their gut contents given
 their higher concentration of prey DNA than that of the cephalothorax
 (Krehenwinkel <i>et al.</i> 2016; Macıas-Hernández <i>et
 al.</i> 2018). To ascertain optimal extraction technique, samples
 were split into two groups. From
 one group, DNA was extracted from the abdomens via Qiagen TissueLyser II
 as per the DNeasy Blood & Tissue Kit (Qiagen, Hilden, Germany)
 manufacturer’s protocol and abdomens kept in the lysis buffer during
 incubation. From the other group, DNA was extracted by splitting the
 abdomen with a sterile micropestle, swilling it in the lysis buffer and
 then removing the bulk tissue. Neither method ultimately
 afforded a significantly greater proportion of prey DNA reads
 post-amplification (Figure S1), so were combined for analysis. Post-lysis,
 all extractions followed the DNeasy Blood & Tissue Kit (Qiagen)
 manufacturer’s protocol but with an extended lysis time of 12 h
 (recommended: 1-3 h) to account for the complex and branched gut system in
 spider abdomens (Krehenwinkel <i>et al.</i> 2016). Per 12
 spiders, each DNA extraction session included at least one negative
 control consisting of an empty tube treated identically to the
 samples. Primers were labelled with unique 10 bp
 molecular identifier tags (MID-tags) and samples had a unique pairing of
 forward and reverse tags for identification of each sample
 post-sequencing. PCR reactions of 25 µl reaction volumes contained 12.5 µl
 Qiagen PCR Multiplex kit, 0.2 µmol (2.5 µl of 2 µM) of each primer, 2.5 µl
 DNase-free water and 5 µl template DNA. Reactions were carried out in the
 same Veriti Thermal Cycler (ThermoFisher Scientific, Waltham, USA), with
 annealing temperatures optimised via temperature gradient PCRs in the same
 machine. PCRs comprised 15 min at 95 °C, followed by 35 cycles of: 95 °C
 for 30 seconds, the primer-specific annealing temperature for 90 seconds,
 and 72 °C for 90 seconds; followed by a final extension at 72 °C for 10
 minutes. The new primers, designated BerenF-LuthienR (universal) and
 TelperionF-LaurelinR (spider-excluding), used annealing temperatures of 52
 °C and 42 °C, respectively. Within each PCR 96-well
 plate, 12 negative (extraction and PCR) and two positive controls were
 included following Taberlet <i>et al.</i> (2018). Negative PCR
 controls consisted of DNase-free water. Positive controls comprised
 known-concentration mixtures of the invertebrate DNA used for primer
 testing, detailed above, quantified using Qubit dsDNA High-sensitivity Assay Kits
 (ThermoFisher Scientific) to ascertain any effects of primer
 bias. All concentrations were standardised at 0.1 ng
 µl<sup>-1</sup> by diluting the DNA in DNase-free water. Five
 mixtures of different species richness and proportions were prepared
 (Table S1). A negative control was present for each MID-tag to identify
 any contamination of primers. Each plate was pooled according to
 concentrations determined by Qiaxcel Advanced System (Qiagen). Each pool
 was cleaned via SPRIselect beads (Beckman Coulter, Brea, USA), with a
 left-side size selection using a 1:1 ratio (retaining ~300-1000 bp
 fragments). The concentration of the pooled DNA was determined via Qubit
 dsDNA High-sensitivity Assay Kits, quality-checked via TapeStation 2200
 (Agilent, Santa Clara, USA) and all pools sharing the same primer pair
 were pooled again into a ‘super pool’, thus forming one pool per primer
 pair. Library preparation for Illumina sequencing was carried out on these
 cleaned ‘super pools’ via NEXTflex Rapid DNA-Seq Kit (Bioo Scientific,
 Austin, USA) and samples were sequenced on an Illumina MiSeq via a Nano
 chip with 2x250 bp paired-end reads (expected capacity ≤1,000,000
 reads). 5.2.5. Bioinformatic analysis
 The Illumina run generated 405,270 and 482,249 reads using
 BerenF-LuthienR (universal) and TelperionF-LaurelinR (spider-excluding),
 respectively. All reads were quality-checked and trimmed in Trimmomatic
 v0.38 (Bolger <i>et al.</i> 2014) with a minimum quality score
 and sliding window of 20 and 4 bp, respectively, and a minimum length of
 135 bp. The read pairs were aligned via FLASH v1.2.11 (Magoč and Salzberg
 2011) and demultiplexed via Mothur v1.39.5 (Schloss <i>et
 al.</i> 2009), removing the MID and primer sequences. Replicates
 were removed, and denoising and clustering to zero-radius OTUs (ZOTUs;
 clustered without % identity to avoid multiple species represented within
 a single OTU) completed via Unoise3 in Usearch11 (Edgar 2010). The
 resultant sequences were assigned a taxonomic identity from GenBank via
 BLASTn v2.7.1. (Camacho <i>et al.</i> 2009) using a 97%
 identity threshold (Alberdi <i>et al.</i> 2017). The BLAST
 output was analysed in MEGAN v6.15.2 (Huson <i>et al.</i>
 2016). Where the top BLAST hit, determined by lowest e-value, was resolved
 at a higher taxonomic level than species-level, the results were checked
 by blasting the sequence manually in GenBank and comparing the results;
 where possibly erroneous entries were preventing species-level assignment
 (e.g. poorly-resolved identifications on GenBank), finer resolution was
 considered. Where ZOTUs were assigned the same taxon, these were
 aggregated. Given the prevalence of family-level assignments (e.g.
 Chloropidae), the data were eventually converted to family-level, but were
 retained at their respective output assignments for clean-up.
 To clean data prior to statistical analysis, all read counts less
 than the maximum read count present in blanks (negative controls and
 unused MID-tag combinations) for its respective ZOTU were removed.
 Instances of non-positive control taxa present in positive controls were
 calculated as a percentage of the maximum read count for that taxon. The
 greatest of these percentages was used to guide a universal percentage of
 the maximum read within each taxon to be removed. This accounts for
 tag-jumping and “bleeding” of over-represented taxa into other samples
 during sequencing. For BerenF-LuthienR, 0.54 % was optimal, whilst there
 were no obvious instances for TelperionF-LaurelinR, so the conservative
 0.54 % was also applied for that library.
 Simultaneously, known lab contaminants (e.g. German cockroach
 <i>Blatella germanica</i> (L.) and various species for which
 molecular analysis was recently undertaken that could be differentiated
 from the target taxa in this study, such as tropical species) were
 identified and the percentage of these occurrences of the total read count
 for their respective samples was calculated. The highest of these
 percentages was used as a guide for the universal percentage of each total
 sample read count to be removed. This accounts for environmental and lab
 contamination, and artefacts and errors of the sequencing process, which
 for BerenF-LuthienR and TelperionF-LaurelinR were 0.43 % and 0.45 %,
 respectively. The data from the two libraries were then aggregated
 together, first removing non-target taxa (e.g. fungi) and instances in
 which predator DNA was amplified (i.e. ZOTUs matching the individual
 spider’s morphological identity). All taxa were converted to family-level
 to standardise the taxonomic level since many ZOTUs could not be resolved
 further; this also increases evenness for subsequent analyses. Whilst all
 conspecific reads were removed to account for predator amplification,
 interspecific linyphiid interactions were still retained, thus any counts
 of linyphiids in the diet exclusively represent the consumption of other
 species. Finally, read counts were converted to presence-absence
 data.
description: Money spider dietary choice in pre- and post-harvest cereal
 crops using metabarcoding - Dryad dataset
 <strong>TelperionFLaurelinRflushcrush.csv</strong>
 gives the percentage of each sample comprised of prey reads for the two
 extraction methods employed: "crushing" which involved beat
 beating of spider abdomens and retention of all tissue for lysis, and
 "flushing" which involved splitting the abdomens, swishing them
 around the tube and removing them prior to lysis.
 <strong>BerenFLuthienRflushcrush.csv</strong> gives
 the percentage of each sample comprised of prey reads for the two
 extraction methods employed: "crushing" which involved beat
 beating of spider abdomens and retention of all tissue for lysis, and
 "flushing" which involved splitting the abdomens, swishing them
 around the tube and removing them prior to lysis.
 <strong>InvertData.csv</strong> presents the
 invertebrate community data of those invertebrates collected by vacuum
 sampling. Abundances are absolute. The ENNR code corresponds to the
 grouping for the EcoNullNetR analyses, based upon pre- and post-harvest
 and the two adjacent fields sampled.
 <strong>DietaryData.csv</strong> presents the data
 derived from HTS of spider gut contents as presence/absence values. As
 with the InvertData.csv, the ENNR code corresponds to the grouping for the
 EcoNullNetR analyses, based upon pre- and post-harvest and the two
 adjacent fields sampled.
 <strong>ENNRDietData.csv</strong> gives the dietary
 data presented in a format compatible with EcoNullNetR.
 <strong>ENNRInvertData.csv</strong> gives the
 invertebrate community data presented in a format compatible with
 EcoNullNetR.
 <strong>ENNRDiet.fl.csv</strong> gives the allowed
 links for the EcoNullNetR analysis.
 <strong>SpiderDietSummary.xlsx</strong> gives the
 dietary data summarised as percentages for each prey taxon and
 grouping.
 <strong>PrimerMinerInSilicoResults.csv</strong> gives
 the percentage of each taxonomic database that was deemed amplifiable by
 each primer pair (with their differences indicating bias).
 <strong>Mock Community Mix 1-5.csv</strong> are five
 files, each denoting the proportion of reads for each taxon with the two
 primer pairs, and the expected proportion given the proportions used in
 the mixes.
 <strong>HarvestMoneySpiderDiet.R</strong> is the R
 script used to generate these data.
 <strong>README.csv</strong> contains these
 descriptions.