Adaptation and integration of remote setting, selective breeding and triploid production technologies to revitalize oyster culture in Delaware Bay

Final Report for FNE11-716

Project Type: Farmer
Funds awarded in 2011: $15,000.00
Projected End Date: 12/31/2012
Region: Northeast
State: New Jersey
Project Leader:
Thomas Foca
Harbor House Seafood, LLC
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Project Information

Summary:

The purpose of this project was to demonstrate the potential use of triploid disease-resistant spatted shell as a production method to revitalize oyster production on leased oyster grounds in lower Delaware Bay. We constructed a remote setting operation, and set eyed larvae on shell from our shucking operation. Setting rates varied due to the quality of the larvae, but rates as high as 12-14 percent, which approximates average levels in the literature were obtained. A total of 420 bushels of spatted shell containing nearly 2 million spat were planted. Survival going into winter was 40% and the estimated survival to market could produce over 668 bushels of marketable oysters. These estimates are optimistic as we were not able to determine the very early mortality of spat smaller than 4 mm. The project initially planned to produce the spat on shell before the growing season, but delays beyond our control and problems with hatchery production forced the project to be conducted later in the year. Achieving this earlier production will likely improve results significantly and reduce time to market by 6 to 12 months.

Introduction:

Farm Profile

Harbor House Seafood is headquartered in Seaford, DE where it operates an oyster packing facility as well as a wholesale and retail seafood market. It operates a fleet of trucks delivering throughout the country including daily air shipments. In October 2007, it acquired facilities in Port Norris, NJ, which is situated directly on the Delaware Bay with its own fleet of boats. This facility, formerly known as the Peterson Packing House, is the last remaining oyster shucking operation in New Jersey. Both facilities are HACCP approved and consistently pass quarterly inspections. Harbor House Seafood, Inc uses its fleet of boats to harvest oysters from public beds in season and its own private beds throughout the year. In Delaware Bay, Harbor House holds 647 acres of oyster grounds leased from the state of New Jersey (section A lots 91, 99, 108 and 153; section B lots 137, 138, 278, and 279; section C lots 35, 109 and 516) and possesses an Aquatic Farmer License from the state of New Jersey to cultivate oysters on its leases in Delaware Bay. For this project, the 58 ft wooden hull Martha Meerwald oyster boat with dual side dredges (Figure 1) was used to access and sample the research sites. Additional vessels in Port Norris include the 72 ft steel hull 44 ton oyster boat “Shell Game II” (also with dual side dredges), the 40 ft Janet R with a stern mounted dredge, and a 24 ft Carolina Skiff. All dredges are commercial 50-inch dredges. All vessels have radar, GPS with SeaMap navigation that can log coordinates and are computer compatible. Two Case frontend loaders (860 and 1400) two Grove cranes (18 ton and 5 ton capacity), and an 80 x 2 ft loading conveyor belt are available to move materials and equipment around property and onto fishing vessels. The Port Norris facility includes an acre of property along the Maurice River (150 ft of river footage) with a 3,000 square ft building and a commercial dock with 100 ft of frontage.

Participants

Dr. David Bushek, Associate Professor of Marine and Coastal Sciences at Rutgers University served as technical advisor to the project. Dr. Bushek is a shellfish biologist specializing in the pathology, ecology and culture of commercially harvested shellfish, particularly clams and oysters. His role was to advise in project design and execution, including construction of setting tanks and setting protocols as well as sampling field plantings to evaluate success. He also provided the direct connection with the Rutgers Cape Shore Oyster Hatchery and Aquaculture Innovation Center via the facilities manager, Mr. Greg DeBrosse who produced the eyed triploid disease resistant oyster larvae for this project.

Project Objectives:

The goal of our project was to evaluate the cultivation potential of remotely set triploid disease resistant oysters in on bottom culture across leased grounds of Delaware Bay. Six tasks were pursued toward this goal:

Task 1. Presample and prepare leased bottom ground by scraping to remove siltation, then plant a base layer of shell.

Task 2. Construct remote setting facility to set triploid, disease-resistant larvae on oyster cultch.

Task 3. Set oyster larvae on oyster shell (cultch) and deploy on leased oyster ground.

Task 4. Plant unspatted shell for comparison of natural recruitment with spatted shell planting.

Task 5. Monitor growth & survival of shell plantings as well as natural recruitment on a control area that was cleaned of sediment, but not planted.

Task 6. Analyze results to assess performance.

Cooperators

Click linked name(s) to expand
  • Dr. David Bushek

Research

Materials and methods:

Task 1. During July 2011, Harbor House scraped, cleaned and sampled Delaware Bay leased ground 138C. The ground was effectively void of oysters. The upper end of the lease was good clean bottom with a decent shell base, so it was decided not to plant shell on the lease. The portion with good clean bottom was divided into two plots: one that would receive spat on shell and one that was left alone to catch wild set for comparison.

Task 2. The Port Norris facility includes an acre of property along the Maurice River (150 ft of river footage) with a 3,000 square ft building and a commercial dock with 100 ft of frontage. The building had been used for storage and was not functional for this project. It was cleaned out, painted and 120 sq ft converted into laboratory space for this project (Figure 2). The laboratory portion was air conditioned and included a 10 ft lab bench. Laboratory equipment included a dissecting scope to count and assess larval viability and spat abundance, a refractometer to determine salinity, a refrigerator to hold ALGAL paste (concentrated phytoplankton) to feed oysters during setting, a water quality test kit, and an assortment of buckets, beakers and graduated cylinders for handling larvae.

Ground on the property was leveled to create a foundation to hold three 3,000 gallon fiberglass tanks (53” deep x 130” in diameter) for setting oyster larvae (Figure 3). Additional space was cleared to accommodate a total of 12 tanks for future expansion. The tanks were plumbed to receive salt water from the adjacent Maurice River as independent static or flow through systems. A 3 hp 3 phase seawater pump was installed inside the building and 2” pvc used to draw seawater from the Maurice River to supply the tanks (Figure 4). Tanks were fitted with 25-micron bag filters to remove zooplankton present in the river water that may act as predators or competitors during setting of oyster larvae (Figure 5). A 2.5 hp 3 phase regenerative blower (Figure 6) was installed in the building to provide aeration via 2” PVC pipe laid in a double ‘H’ configuration along the bottom of the tanks (Figure 7). Holes drilled into the top of the pipes were variously plugged with stainless steel screws until air flow was balanced across the tank when filled with water. A triple tube straight immersion heaters was coupled to a heat controller and mounted on the side of one tank to allow for early season production by increasing ambient water temperature to 25oC to promote setting of eyed larvae (Figure 8).

Oyster cultch (shucked oyster shells) were accumulated during the previous year’s shucking operation and stored in open air to cure for this project. Shells were pressure washed with fresh water to remove debris prior to use. To contain the shell in the setting tanks and facilitate moving shell from the shucking house to the tanks and from the tanks to the Martha Meerwald for planting, 30 pvc coated heavy wire metal cages were constructed to hold approximately 20 bushels of oyster shell each (Figure 9). Cages consisted of two rows of four columns separated by a 4” channel. This design follows that developed by the University of Maryland, except their design included a second channel bisecting the rows in half. By eliminating this cross channel we increased the capacity of shell that could be added to a setting tank by allowing us to place eight cages in a tank instead of six. We tested whether or not this decreased larval access to the shell (see results), as this was the primary reason for the channels.

Task 3. An Case frontend loader and an 80 x 2 ft loading conveyor belt were used to fill cages with oyster shell that had been retained for this project by the Harbor House shucking operation during the previous year. A 5 ton Grove cranes was used to move cages filled with oyster shell into and out of setting tanks and onto the deck of the Martha Meerwald for planting on our oyster lease (Figure 10a and b).

Disease resistant triploid eyed larvae (i.e., mature larvae ready to metamorphose and attach to oyster shell) were received on July 30th (10.9 million), August 5th (9.7 million), and August 16 (2.6 million). These dates were later than originally planned due to construction delays and subsequent hatchery delays but corresponded to the period of natural oyster recruitment in Delaware Bay enabling us to continue the project as planned. In total, 23 million eyed larvae were purchased from the NJ AIC for use in this project. Upon arrival, larvae were inspected with Dr. Bushek instructing our staff how to handle and evaluate them for viability and abundance, and then prepare them for adding to the setting tanks. Eight cages were added to a setting tank for each batch of larvae two to three days before delivery of eyed larvae to allow natural microbial films to develop that encourage setting of oyster larvae. Tanks were filled with 25 micron filtered river water during high tide (slack high plus up to one hour past slack) to obtain the highest salinity water with the least sediment. Crystal Sea Salt(registered trade mark) was added as needed to increase salinity to 20 ppt to minimize differences between hatchery conditions and setting conditions. Salinity was monitored regularly before and during setting. The eyed larvae were then resuspended in a five gallon bucket containing about four gallons of water from the setting tank (Figure 11). Larvae were continuously mixed with the bottom of a graduated cylinder and inspected for activity, then added to the setting tank by pouring about half a liter at a time into different portions of the tank to ensure larvae were well dispersed (Figure 12). Tanks were left static after larvae were added for a period of three days to ensure larvae time to set (i.e., attached to oyster cultch).

To test the effect of removing the second cross channel in the shell cages, two central columns were left empty in four of the eight cages such that each column had at least three exposed sides (Figure 13). The four full cages contained four columns with only two exposed sides, possibly limiting access by larvae. Because differences were not detected during the first or second setting trial, all columns were filled for the third trial.

After adding larvae, 500 ml of concentrated phytoplankton (Instant Algae Shellfish Diet (registered trade mark), Reed Mariculture, Cambell, CA) was added to the tank and repeated twice daily while the tank was static and larvae were setting. Subsequently, feeding rations were increased to two liters per day as they larvae and spat were clearing the water column in the tank. On day three, the seawater pump was turned on to provided a continuous supply of food from the River. During this process, the salinity of the tanks equilibrated to that of the river water as the water in the tanks was replaced with river water. Shells were haphazardly collected on day two for the presence of spat (Figure 14). Spatted shells were planted on August 8th, August 15th, and on August 26, 2011 (Figure 15). The third planting was performed two days before hurricane Irene hit the coast of South Jersey. Before each planting shells were gathered from the top, middle and bottom of each cage to determine setting efficiency (that is, the percentage of the eyed larvae that had successfully attached to shells being planted).

Task 4. Because the bottom of the leased ground contained an adequate shell base following cleaning, no shell was planted.

Task 5. Our initial plan was to set larvae as soon as possible in May or June to extend the initial growing season so that oysters would get a head start on reaching predator size refuges and also reach market size earlier. Unfortunately, delays in construction and, more importantly, delays in hatchery production beyond our control that resulted from an unusual coast-wide larval mortality event, did not permit this early setting of oyster larvae. Nevertheless, we did successfully set larvae later in the year. The first planting was not conducted until August 8th and the third occurred on August 26, 2011. Two days after the third planting hurricane Irene hit the coast of South Jersey. Hurricane Irene was followed by Tropical Storm Lee and the combined effects of these storms caused unprecedented fresh water inflow (flooding) into the Delaware Bay. This dilemma led to many detrimental environmental conditions that threatened the survival of our newly planted spat on shell and detracted from our ability to completed monthly monitoring. Our science advisor from Rutgers, Dr. David Bushek, was able to deploy a salinity recorder directly over our shell plant within a week of the storms passing and determined that salinity levels recovered quickly although remained on the low side of average levels through September as freshwater worked its way down the bay. As a result, samples were only collected in November 2011.

Task 6. Results were analyzed by calculating setting rates (the percentage of eyed larvae that ended up on shell to be planted), estimating growth rate of oysters once planted and estimating survival through November 2011.

During the first and second remote setting we tested whether or not eliminating the cross channel in the shell container restricted access to shell by the setting oyster larvae. This was accomplished by leaving 2 center columns empty on 4 of the eight cages. At the time of the first planting, 25 shells were removed from each cage after it was dumped on board the Martha Meerwald (Figure 16). Spat were counted under a stereoscope in the laboratory and 25 measured to determine size at deployment. During the second planting, care was taken to remove shells separately from the top, middle and bottom of each cage (5 from each section) as the cage was emptied on the deck of the Martha Meerwald for transport to the leased ground and planting. For the third planting, 55 shells were collected from what was deposited on the deck of the Meewald to plant. Each group of shells was labeled and returned to the laboratory to be counted and measured (Figure 17). To evaluate on bottom survival and growth, a systematic sampling was accomplished in November 15, 2011 using Rutgers Haskin Shellfish Laboratory R/V Veliger and a small 0.81 m oyster dredge. Several tows were taken to obtain 45 shells with spat over the planted portion of the lease. No shells with spat were detected elsewhere on the leased ground used for this project.

Research results and discussion:
Results of remote setting

The three separate remote settings generated about 2 million spat on shell for planting: approximately 1.5 million spat on 130 bu from the first planting, 0.4 million spat on 130 bu for the second planting and 0.1 million spat on 160 bu for the third planting. In all 420 bushels of triploid, disease-resistant spatted shell were planted demonstrating the potential of this technique if scaled properly.

The first setting was performed with 10.9 million eyed larvae. On day 2, 15 shells were gathered haphazardly from around the tank and examined under a microscope for spat. All but three shells contained spat, the most with 92, and the average was 21. Counts of shells added to columns revealed that each column holds approximately 1,275 shells so the total number of spat at that point was estimated at 1.39 million or about 13% of the larvae that were added to the tank. Estimates of spat abundance collected on day 10 as shell were loaded for planting confirmed a total of 1.4 million spat from the 10.9 million larvae added to the tank for a final setting rate of 12.8%. This is about average when compared with published reports.

The second setting was not nearly as successful. The 9.7 million larvae were much less responsive when resuspended and many appeared dead. Because we were nearing the end of the spawning season we decided to continue the work as no other sources of larvae were available. Evaluation of setting of this group indicated a total of about 421,590 larvae set successfully and survived to be planted for a setting rate of 4.3%.

The third setting was a last ditch effort to recover what we could from the final batch of larvae at the hatchery. These larvae we considered runts by the hatchery technician. They had not progressed well in the hatchery. Upon receipt, they responded well and many were eyed, but others were not. Of the 2.5 million larvae received, only 26,017 were estimated to have set successfully at the time of planting. This is a setting rate of only 1% and is not acceptable, but was all that we could recover from a difficulty hatchery production year.

Effect of removing cross channel in cages

Oyster set was compared on full cages and cages with two center columns empty to assess the need to increase access of larvae to shell. During the first setting, shells in full cages contained an average of 17.6 spat per shell with a standard deviation of 20.4. The maximum number of spat on a shell was 106 and the minimum was zero. The most common count (i.e., the mode) was 4 and the median was 10.5 indicating that half the shells contained more than 10 spat each. By comparison, shells in partial cages contained an average of 22.5 spat per shell with a standard deviation of 31.5. The maximum number of spat on a shell in a partial cage was 151 and the minimum was zero. The most common count (i.e., the mode) was 6 and the median was 10, indicating that half the shells contained more than 10 spat each. A t-test indicated no significant difference (p = 0.194) between the number of spat on shells in either partial or full cages. Therefore, full cages averaged 22,312 spat per column or 178,500 spat per cage, whereas partial cages averaged 28,687 or only 172,125 spat per cage. The lower density per shell means fewer oysters will grow together and compete for space in the full cages versus the partial cages. Furthermore, the total obtained in the full cages was slightly larger owing to the fact that there was 25% more shell with essentially an equal density of spat.

The absence of a difference was further supported by the second setting trial. Here we compared the density of spat/shell on the top third, middle third and bottom third of each type of cage. This time densities were lower on the partially filled cages: 4.4 spat per shell verus 7 spat per shell. This difference was due almost exclusively to the different setting rates on the upper third of the cages. Spat per shell in the upper third of partially full cages was only 1.9 while those on full cages was 7.5. As a result, full cages contained an average estimate of 71,456 spat whereas partial cages only contained an estimated 33,942 spat. These results combined with those from the first trial indicated that removing the perpendicular cross channel did not affect setting rates. Therefore, the final trial was completed with full cages only.

Survival of spat on planted shell

The November evaluation estimated survival at 40%. This is relatively low and attributed to the impacts of Hurricane Irene and Tropical Storm Lee, which undoubtedly increased sedimentation and then reduced salinity to points that may have increased mortality. Furthermore, this number may be an overestimate as spat smaller than 4.1 mm that may have died were not detected: spat were 1.25 – 6.05 mm when deployed, whereas dead oysters that were sampled on November 15th averaged 8.4 mm in size with a range of 4.1 to 18.8 mm. Surviving oysters averaged 15.9 mm with a range of 7.3 – 39.5 mm, indicating that they may have reached a size refuge from early mortality. This mean size is within the normal range expected for oysters that would have recruited naturally. The earlier production scheduled that was planned would have likely produced larger oysters and increased survival.

Economics

Because this project was designed to demonstrate the potential for producing and deploying triploid disease-resistant spat on shell in lower Delaware Bay, no economic analysis was performed per se. However, even with the poor larval production and setting rates, we can calculate a potential return from the shell plantings. A total of 1.9 million spat on shell were planted by the end of August. Survival was estimated at 40% in November, leaving approximately 742,458 oysters. Rutgers Haskin Shellfish Research Laboratory estimates an average mortality of 35-40% per year for oysters on the lower seedbeds adjacent to the leased grounds. Using 40% per year, there will be 445,475 oysters remaining by Fall 2012, but they will likely be too small to harvest because of the delays in 2011 that missed most of the growing season. Adding another season of growth and mortality will lose another 40% resulting in a final tally of 267,285 oysters. Assuming a harvest success of 75%, Harbor House should be able to recover 200,464 oysters. Assuming 300 oysters per bushel that’s 688 bushels. Assuming a conservative price of $40 per bushel that’s $26,728, making the prospects of this investment within reason, even during a poor year.

Conditions

Clearly, refined hatchery performance is needed to make the most of this process. Delays in construction along with problems at the hatchery limited the success, but 2011 was an unusual year with reports of poor hatchery survival prevalent along the east coast. Unusual tropical storm activity also created problems that affected results and most likely increased mortality during the fall of 2011. Obtaining eyed larvae in April or May will likely have a dramatic impact on the success of planting triploid disease resistant spat on shell.

Participation Summary

Education & Outreach Activities and Participation Summary

Participation Summary

Education/outreach description:

Two specific efforts in outreach were performed concerning this project. First, we created a power point presentation to explain the project and presented it during an oyster forum organized by the Bayshore Discovery Project located in Bivalve, NJ. A copy of this presentation is uploaded as a pdf file. Second, we set up a display at the Bay Day Festival in Bivalve, NJ that documented the process and staffed a booth to explain the project to festival attendees.

Project Outcomes

Assessment of Project Approach and Areas of Further Study:

Future Recommendations

Adoption

Plans were underway to double the effort this past year and produce spat earlier in the season as a follow up study. Unfortunately, the storms from last year and the derecho that came through our region in late July 2012 as well as other developments have redirected our efforts for the moment. Once we have recovered from these issues we plan to repeat the study with spat produced earlier in the year.

Assessment

The primary problem that we need to overcome is obtaining larvae earlier in the season. We have identified at least one alternative producer and have leads on several others that could help fulfill our needs. In terms of monitoring the project, it would be useful to conduct the test plantings using shell other than oyster shell. Surfclam or ocean quahog shell is available and commonly used to replant oyster beds. It provides a more easily traced substrate as these shells are not naturally present in the bay so anything attached to them would clearly be from our plantings.

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.