Methods to control bio-fouling of cultured eastern oysters, Crassostrea virginica, by the tube-building polychaete worm, Polydora cornuta

Project Overview

FNE13-780
Project Type: Farmer
Funds awarded in 2013: $13,415.00
Projected End Date: 12/31/2014
Region: Northeast
State: New Jersey
Project Leader:
Betsy Haskin
Email
Betsy's Cape Shore Salts

Annual Reports

Commodities

  • Animals: shellfish

Practices

  • Animal Production: general animal production
  • Education and Training: extension, farmer to farmer, on-farm/ranch research
  • Pest Management: chemical control, physical control

    Proposal summary:

    The purpose of this project is to develop efficient and effective methods for control of Polydora infestation in eastern oysters. Two control methods, hypersaline dips and lime dips, which have been used with success to control similar infestations in Pacific oysters grown in Mexico and Australia, will be evaluated for efficacy in controlling Polydora infestations at an oyster farm in Delaware Bay, NJ. Life history aspects of the worm, including season planktonic abundance and settlement patterns, will be investigated to establish treatment regimes that target the worm at the onset of infestation. Results will be presented locally at the Rutgers University hosted Delaware Bay Shellfish Growers Forum and regionally at the Milford Aquaculture Seminar. Web publishing on eXtension will occur if the results prove beneficial. Oyster farms in the northeastern US and worldwide are plagued by biofouling caused by marine polychaetes belonging to the genus Polydora. Two species are problematic in the northeast of the US, Polydora websteri and Polydora cornuta (formerly ligni). Generally referred to as mud-worms, these ubiquitous polydorid worms can kill oysters, reduce oyster growth, fragment the oyster’s shells making them difficult to shuck, and cause internal shell blisters that decrease product appeal and marketability. Oyster farm profitability is limited by mud-worm associated oyster mortalities and by the extensive time and effort invested in mud-worm control. The issue has existed as long as farmers have been cultivating oysters; however, mitigating treatments have been met with limited success. Location, local conditions, and cultivation methods play an important role in modulating infestation timing and severity, and the efficacy of control methods. Nationally, oyster farming is a 300+ million dollar industry supporting thousands of small farms and sustainable green jobs. Farm-raised oyster production has increased rapidly in the Northeast and mid-Atlantic region during the last decade. In New Jersey production is expanding as a result of new state leasing policies and will exceed one million oysters this year; production is anticipated to exceed five million oysters in the next five years. Cultivated single oysters yield $0.50 to $0.85 wholesale. New Jersey’s oyster farms are concentrated on the extensive intertidal sand flats of the lower Delaware Bay (Cape Shore) where they are exposed twice daily during low tide. The water with its moderately high salinity and rich food quality supports rapid oyster growth and yields exceptional quality oysters. Here hatchery reared oysters are grown in plastic mesh bags that are secured to rebar frames positioned on the bay bottom. The farms are accessed from the shore at low tide and the bags are tended for a 1-3 year production cycle. Oyster culture systems provide ideal substrate for Polydora ettlement and growth. During the spring and summer oyster farmers must clean the worms and their thick mud tubes from the oyster bags in order to avoid oyster mortality caused by the mud smothering the oysters. These mud tubes were found to cause extensive oyster mortalities in Delaware Bay by Stauber and Nelson (1940) who found as many as 36 worms per cubic centimeter. The dry weight of mud accumulated by these worms has been estimated to be 98 tons per acre (Orth,1971). The process of washing the mud and worms from the oyster bags requires significant effort and time, about 700 man-hours annually for a midsize farm producing 250,000 oysters. When combined with equipment and supply costs Polydora control costs are significant. In a recent Shellfish Aquaculture Needs Assessment conducted by Rutgers University control of fouling organisms was ranked as a high priority issue by 90 the respondents. The development of more effective and efficient control measures are critical to reducing labor cost, improving yields and oyster quality, and enhancing farm profits.

    Project objectives from proposal:

    The purpose of this project is to develop a better understanding of the life history and biology of the polycheate worm Polydora cornuta and to develop efficient and effective methods for control of its infestation to the oysters.

    Specific objectives are:
    1. Conduct plankton and settlement surveys to determine temporal variability in abundances of Polydora cornuta larvae and peak settlement periods to inform and improve the efficacy of treatment timing and options.

    2. To evaluate the effectiveness of time targeted hypersaline and lime dip treatments as a control for P. cornuta. Polydora Field

    Survey (Objectives 1)

    The study will be conducted at my farm located in on the Cape Shore flats of the lower Delaware Bay at Green Creek. Weekly larvae samples will be collected with a Plankton net (100 micron mesh) beginning in early April 2013 and continuing through August 2013. Thereafter, monthly samples will be collected through October 2013.

    Surface tows will be collected on falling tide approximately two hours before low tide. Plankton samples will be processed as described by Orth (1971). Briefly, samples will be subsampled with a plankton splitter and Polydora cornuta larvae will be identified and counted. Subsamples will be preserved for later measurements of setiger length using an ocular micrometer.

    In order to assess the timing of larval settlement, triplicate shell bags containing 8 liters of clean oyster shells will be placed on rebar racks at the farm in March 2013. Five shells from each bag will be removed weekly from April through October and examined for Polydora in order to determine settlement periods. Additionally shell strings(n=3) containing 5 shells each will be deployed for one-week intervals from March through October and similarly examined for newly recruited worms. Prior to examination the shells will be incubated in seawater baths containing 100 ppm o-dichlorobenzene for six hours to remove worms from shell crevices (MacKenzie and Shearer, 1959). Total numbers of P. cornuta will be enumerated and size measurements will be taken on a subsample of 20 worms. Examinations will be made using a dissecting microscope. When necessary worms will be preserved in formalin solution prior to quantification. Supporting laboratory equipment and space will be provided by the Haskin Shellfish Research Laboratory of Rutgers University.

    Water temperature at the site will be continuously monitored at 15 min intervals through the course of the study using an Onset data logger. Polydora abundances in plankton and on shell bags and strings will be Polydora field surveys will be conducted in collaboration with scientist Rose Petrecca, Institute of Marine and Coastal Sciences and Lisa Calvo, Program Coordinator I, Haskin Shellfish Research Laboratory. An undergraduate student will be hired to assist with the project.

    Polydora Control Evaluations

    Two immersion dip treatments(hypersaline and lime) will be evaluated for efficacy in reducing P. cornuta fouling relative to standard seawater water spray washing. Fifteen oyster bags containing an eight-liter volume of one year old oysters will be randomly assigned to one of three treatments—hypersaline dip, lime dip, or seawater wash. The five replicate bags for each treatment will be labeled will be deployed on the farm in a block design with 1-replicate from each secured on each of five racks whose location on the farmed will be randomly assigned.

    Treatment bag placement on each of the racks will also be assigned randomly. Immersion solutions, lime (0.2% calcium hydroxide, 2 g L-1) and hypersaline (200 ppt salt) will be prepared in 40-gallon cans using ambient seawater. Oyster bags will be removed from the rebar racks and placed vertically in the bath for 5 minutes for lime treatment and 15 minutes for hypersaline treatment. After immersion the bags will be allowed to dry on the rebar racks for a minimum of 30 minutes. Drying time and temperature of treatment baths will be noted. At the same time bags assigned to seawater spray washing will be treated using a Honda gasoline powered water pump and hose.

    Treatments will be applied as soon as plankton and settlement samples
    indicate Polydora larvae are present and have begun to settle. A subsequent treatment will be administered 14 days after the initial treatment and as needed through the remainder of the summer. Polydora infestation of treatment bags will be measured via visual inspection of the entire bag and ranked as negative, light, moderate, and heavy each week. Twice monthly, three oysters will be removed from each treatment bag and quantitative evaluations of the number of Polydora worms will be conducted following the methods described above for settlement on shell bags.

    Oyster mortality and growth rates will be assessed in July and September. On each date all dead and live oysters in each treatment bag will counted and the shell heights of a subsample of 50 live oysters will be measured using digital calipers.

    Statistical analyses of fouling ranks, oyster mortality, growth (shell height), and worm abundance will be analyzed using ANOVA tests on appropriately transformed data.
    References follow:

    Mackenzie, C. and L. Shearer. 1959. Chemical control of Polydora websteri and other annelids inhabiting oyster shells. Proc. Natl. Shellfish Assoc 50:105-111

    Orth, R. 1971. Observations on the Plantonic Larvae of Polydora ligni Webster (Polychaeta:n Spioniae) in the York river, Virginia. Chesapeake Science Vol. 12, No. 3, p.121-124.

    Any opinions, findings, conclusions, or recommendations expressed in this publication are those of the author(s) and do not necessarily reflect the view of the U.S. Department of Agriculture or SARE.