Dataset: Oyster genetic identity and parasite community structure

Oyster Reef Restoration Experiment
In May 2017, we created nine oyster reefs in Quonochontaug Pond, Rhode Island (RI) (41°20'22.6"N, 71°43'41.2"W), with three separate reefs each located within three distinct regions (West, Northeast, East). Each ~22 square meter (m²) reef (0.5-0.8 m height) was constructed from a base layer of steam-shucked clam shell topped with clean, recycled oyster shell, and then seeded with remote-set spat on shell (see Davenport et al. 2022 for a detailed description of reef construction). A goal of this study was to compare the performance of different oyster sources and the consequent structure of associated parasite communities, so one replicate reef per oyster source was constructed in each of the three regions of Quonochontaug Pond (hereafter, blocks). The oyster seed sources included one line from a regional commercial hatchery, as well as two wild progenitor lines spawned from broodstock collected from nearby existing wild populations in Green Hill Pond, RI and Narrow River, RI. All oyster lines were spawned at local hatcheries, transferred to Roger Williams University Shellfish Hatchery and set on oyster shell in June 2016, and then stored in cages on an oyster lease in Quonochontaug Pond until reef construction in May 2017.

Reef Monitoring and Oyster Collection
In the fall of each year for four years post-restoration (2017-2020), we monitored oyster density and size distribution by non-destructively sampling six haphazard 0.25 m² quadrats per reef and recording the number of live and dead oysters, as well as shell height of a subsample of up to 50 live and 30 dead oysters (following methods of Griffin et al. 2012 (see Supplemental Files) and guidance of Baggett et al. 2015). Coincident with each fall monitoring event, we harvested ~35 haphazardly-selected live oysters from each reef for analysis of parasite communities; samples were transported to the Northeastern University Marine Science Center on ice and then stored at -80 degrees Celsius prior to processing.

Parasite Prevalence and Intensity
Crassostrea virginica is commonly infected by a variety of micro- and macro-parasites simultaneously. We assessed the prevalence (proportion of sampled oysters infected per oyster reef) and intensity (parasite concentration per infected host) of five common parasite species (microparasites: Perkinsus marinus (Dermo disease), Haplosporidium nelsoni (MSX disease), and Haplosporidium costale (SSO disease); macroparasites: Cliona spp. and Polydoraspp.).

To assess microparasite prevalence and intensity, DNA was extracted from up to 32 oysters per reef and then amplified using both a polymerase chain reaction (PCR) assay modified from Stokes & Burreson (2001) SSO protocol, and a multiplex quantitative polymerase chain reaction (qPCR) assay modified from De Faveri et al. (2009) Dermo protocol and Wilbur et al. (2012) MSX protocol. DNA was extracted using 20-40 milligrams (wet weight) of gill and mantle tissue with the Omega Bio-Tek E-Z 96® Tissue DNA Kit.

We used a modified version of the De Faveri et al. (2009) Dermo protocol and the Wilbur et al. (2012) MSX protocol to analyze the samples on a Bio-Rad CFX96TM Real-Time System with Bio-Rad CFX Manager Software (version 3.1). Each reaction consisted of 1 microliter (μl) template DNA, 3 μl water, 5 μl TaqMan Multiplex Master Mix, and 0.5 μl of each 20X primer/probe master mix, which contained 18 μM of each P. marinus primer and 5 μM of TaqManTM QSY probe (De Faveri et al. 2009) for Dermo, and 18 μM of each H. nelsoni primer and 5 μM of TaqManTM MGB probe (Wilbur et al. 2012) for MSX. qPCR cycling conditions included initial denaturation at 95°C for 30 sec, followed by 40 cycles of 95°C for 10 sec and 60°C for 30 sec. We used gBlocks® (gene fragments containing the target regions from P. marinus and H. nelsoni; Integrated DNA Technologies) to develop standard curves, and extracted DNA from cultures of P. marinus and H. nelsoni with known cell quantities to use as positive controls. All standards, samples, and positive and negative controls were run in duplicate; if samples differed by >1 Cq, they were rerun to confirm presence/absence and/or parasite concentration.

We used a modified version of the Stokes & Burreson (2001) SSO protocol to assess H. costale prevalence on a Bio-Rad CT100 thermal cycler. PCR cycling conditions included initial denaturation at 94°C for 2 min, followed by 35 cycles of 94°C for 30 sec, 59°C for 30 sec, and 72°C for 60 sec, and final extension at 72°C for 5 min. PCR products were visualized on a 1% agarose gel and photographed for analysis of band intensity using ImageJ, following the methods of DeLong & Hanley (2013). For the SSO assay, we used gBlocks® containing the target region from H. costale as positive controls and as standards for estimating parasite concentration based on quantification of band intensity using ImageJ (Abràmoff et al. 2004).

To assess the prevalence and intensity of two common macroparasites, boring sponge (Cliona spp.) and mud blister worm (Polydora spp.), we photographed the inside (mud blisters) and/or outside (sponge holes) of both top and bottom valves for all samples with holes characteristic of boring sponges and blisters characteristic of mud blister worms to quantify the proportion of affected shell area (i.e., [(total infected area/total shell area)*100]) using ImageJ (Abràmoff et al. 2004), following the methods of Hanley et al. (2019).