Abstract:

Oyster populations, dermo disease, epifaunal communities and water quality were monitored for three years (2020-06-05 to 2024-02-08) on an area of newly restored reefs at Grass Islands Reef, Aransas Bay, Texas. These biotic variables were monitored on high relief "mounds", low relief "flats" and unrestored control areas. Monitoring occurred immediately prior to restoration, every month for three months after construction, and quarterly thereafter. The purpose of the monitoring was to compare oyster populations and associated epifaunal communities on the two restored reef types.

Suggested Citation:

Palmer, T.A., M.S. Sugla, D. White, N.J. Breaux, and J. Beseres Pollack. 2024. Field observations of oyster populations, dermo disease, and epifaunal communities on high- and low-vertical relief restored oyster reefs at Grass Islands Reef, Aransas Bay, Texas from 2020-06-05 to 2024-02-08. Distributed by: GRIIDC, Harte Research Institute, Texas A&M University–Corpus Christi. doi:10.7266/dqnde5q5

Purpose:

The purpose of this study was to compare oyster and epifauna densities on mound and reef flats to each other, and to an unrestored control.

Data Parameters and Units:

Date (ddMMMyyyy), Oyster height (mm), temperature (°C), specific conductance (mS/cm), salinity (PSU), dissolved oxygen (% and mg/L), pH, turbidity (NTU), secchi disc (m), live oyster abundance (n), dead oyster abundance (n), volume of live oysters (L), volume of dead oysters (L), volume of rock(L), oyster height (mm), epifauna abundance (n, n m-2), epifauna biomass (g, g m-2), infauna abundance (n, n m-2), infauna biomass (g, g m-2), dermo intensity, (0-5 scale), Total suspended solids (mg/L), ash-free dry weight (mg/L), water chlorophyll (ug/L), dermo intensity (1-5). Blank cells equal no data collected.

Methods:

Pre-construction monitoring was conducted in June 2020 (2 months before reef construction) and August 2020 (0 months). Eleven 0.5 m2 quadrats were sampled by divers in areas selected for construction of reef flats (3 quadrats) and reef mounds (5 quadrats), as well as in areas selected to remain unrestored (3 quadrats) to quantify densities of oyster spat, barnacles, mussels, and live and dead juvenile and adult oysters. The quadrats were excavated to a 5 cm depth, bagged, and assessed for oyster density at the surface. Following reef construction at the beginning of August 2020, nine sampling trays (45 x 30 x 10.5 cm; 0.135 m2) were placed by divers at each of nine sites: three unrestored, three reef mounds, and three reef flats. Each tray contained a single layer of limestone (limestone was the substrate used for the restored reefs also). The trays were secured in place with hooked rebar. One tray was sampled from each of the sites without replacement monthly for the first three months after reef construction (September, October, November 2020), and then quarterly thereafter (February, May, August 2021). Trays were assessed for oyster metrics and motile epifauna. Additional sampling of reef mounds, reef flats, and unrestored areas was conducted using 0.25 m2 or 0.5 m2 quadrats every six months thereafter. Encrusting species were identified to LPIL (usually species) and counted. Shell height was measured for oysters larger than 5 mm. Oysters >25 mm shell height were assessed to determine prevalence and intensity of Perkinsus marinus, the protozoan parasite responsible for Dermo disease. To assess for P. marinus infection, a 5 x 5-mm section of mantle-edge tissue was removed from just over the palps from each oyster and cultured for one week using Ray’s Fluid Thioglycollate culture method (Ray, 1966). Tissues were then stained with Lugol’s iodine solution and examined under a microscope for prevalence and intensity of P. marinus hypnospores. Perkinsus marinus intensity was scored from 0 (uninfected) to 5 (heavily infected) (Mackin, 1962; Craig et al., 1989). Motile reef-resident epifauna (>1 mm) were collected in the field by rinsing the limestone cobble with seawater over a 0.5 mm mesh, and were then placed in 10% buffered formalin. In the laboratory, motile fauna from the trays were sorted, counted, and identified to the lowest practical taxon (usually species). Dry weight biomass for each taxon in each sample was measured after drying organisms for approximately 24 hours at 60°C. Mollusks were placed in 0.1 M HCl to remove shells before weighing. Discrete measurements of the environmental variables (e.g., temperature, dissolved oxygen , and salinity were made at the surface and the bottom of the water column at multiple sampling stations on each sampling date using a YSI ProDSS data sonde. One liter of water was collected by hand in an amber bottle from 0.1 m above the bottom and placed on ice in the dark for analysis of chlorophyll-a, total suspended solids and ash-free dry weight. Water samples were filtered through pre-combusted glass fiber filters (Whatman GF/F glass fiber filters, 0.7 mm porosity). Chlorophyll-a was extracted from filters overnight using a non-acidification technique and read on a Turner Trilogy fluorometer (Turner Designs, Sunnyvale, USA) (Welschmeyer, 1994; EPA method 445.0). Total Suspended Solids (TSS) were measured using EPA method 160.2.

Provenance and Historical References:

Craig, A., Powell, E. N., Fay, R. R., & Brooks, J. M. (1989). Distribution of Perkinsus marinus in Gulf Coast oyster populations. Estuaries, 12(2), 82–91. https://doi.org/10.2307/1351499. Mackin, J.G. (1962). Oyster disease caused by Dermocystidium marinum and other microorganisms in Louisiana. Publ. Inst. Mar. Sci. Univ. Tex. 7:132-299 Ray, S.M. (1966). A review of the culture method for detecting Dermocystidium marinum, with suggested modifications and precautions. Proceedings of the National Shellfisheries Association, 54: 55–69.

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