EAST HAMPTON TOWN SHELLFISH HATCHERY

2016 ANNUAL REPORT AND 2017 OPERATING PLAN

Prepared by Kate Rossi-Snook Edited by Barley Dunne East Hampton Town Shellfish Hatchery

Kate and Adam prepare to seed oysters on a quiet morning in Three Mile Harbor

Annual Report of Operations

Mission Statement

With a hatchery on Fort Pond Bay, a nursery on Three Mile Harbor, and a floating raft field growout system in Napeague Harbor, the East Hampton Town Shellfish Hatchery produces large quantities of oyster (Crassostrea virginica), clam (Mercenaria mercenaria), and bay scallop (Argopecten irradians) seed to enhance valuable shellfish stocks in local waterways. Shellfish are available for harvest by all permitted town residents.

Cooperative research and experimentation concerning shellfish culture, the subsequent success of seed in the wild, and the status of the resource is undertaken and reported upon regularly, often funded and validated by scientific research grants. Educational opportunities afforded by the work include school group and open house tours and educational displays at community functions.

Annual reporting includes production statistics and values, seed dissemination information, results of research initiatives, a summary of outreach efforts, the status of current and developing infrastructure, and a plan for the following year’s operations.

2016

Full-time Staff Part-time and Contractual Volunteers John “Barley” Dunne – Director Adam Younes – Environmental Aide Jannine DeMeritt Kate Rossi-Snook – Hatchery Manager Sam Younes – Environmental Aide (summer) Julia DeMeritt Pete Topping – Algae Culturist Tali Friedman – Environmental Aide (summer) Thor Botero Jeremy Gould – Maintenance Mechanic

Pete checking on the success of our Three Mile Harbor scallop sanctuary

Special Thanks to:

Barnaby Friedman for producing our annual seeding maps

The East Hampton Town Board and East Hampton Trustees for their continued support

Table of Contents

2016 Annual Report of Operations

Season Summary 1

2017 Operating Plan 3

Oyster Production 4 Spawn and Culture Summary Discards and Culls Distribution Overwintering Market Values

Hard Clam Production 6 Spawn and Culture Summary Discards and Culls Distribution Overwintering Market Values

Bay Scallop Production 9 Collection and Culture Summary Discards and Culls Distribution Overwintering Market Values

Projects and Cooperative Research 12 East Hampton Shellfish Education and Enhancement Directive (EHSEED) Montauk Elementary School 8th Grade Science Fair Research: Oyster Murder Mystery Battle of the Bivalves Understanding the effects of predicted ocean conditions as a result of climate change on juvenile shellfish Shinnecock Bay Restoration Project oyster reef construction Impacts of climate change and ocean acidification on economically important shellfish in New York: are there effective mitigation and adaptation measures? Exploring trait-mediated effects of finfish on decapod crustaceans and bay scallops in eelgrass ecosystems Juvenile clam growth and survival in Western Shinnecock Bay

Public Outreach & Industry Involvement 36

Infrastructure Management 38

Appendix I: 2015 Harbor Seeding Maps – All Species 40 II: 2015 Harbor Water Temperatures 48

2016 Season Summary

Oysters: The hatchery season began with an accidental spawn in the broodstock conditioning tank one day before the first scheduled spawn. After conferring with Karen Rivara of Aeros Cultured Oyster Co. and Noank Aquaculture Cooperative, we determined that the usual conditioning time of 8 weeks was too long for the younger-than-usual (2015 product) oysters being used to breed disease-resistant offspring. We salvaged some larvae from the tank, quickly adjusted our spawning schedule, and reintroduced the first oyster cohort back to the conditioning tank to represent a fourth spawn.

For the second consecutive year we experienced some die-off and reduced growth due to Juvenile Oyster Disease/Roseovarius Oyster Disease. The survivors, however, developed well and resulted in an excellent crop. We set aside some of these oysters for spawning in 2017 in order to continue breeding resistance to the disease. Full disease resistance is expected to take several years to develop.

Despite these setbacks we had a robust season of oysters, moving almost 8 million to the nursery, and over 5 million to the field for growout, of which 1.3 million achieved ideal seeding size.

The total value of oyster production in 2016 was $291,552.

Clams: 2016 was a great year for clam production; a total of more than 31 million culls were seeded from the nursery and field, and we produced and disseminated 3.74 million seed clams. We moved a lot of clams to the nursery (nearly 25 million vs. 11.5 in 2015) and produced more seed-sized clams than in 2015. However, the average size of seed clams decreased from 2015 (11.7mm vs. 13.6mm). We will continue to fine-tune clam production in an effort to optimize our “quality vs. quantity” approach.

Unfortunately, overwintered clams did not exhibit much success. Of the more than 1 million clams overwintered in 2015, only about 450,000 were retrieved and seeded (44% survival), and we saw a mere 1% growth. We continued to overwinter clams in Northwest Creek for the 2016-2017 season, but are also trying three other test sites to determine if another site is more suitable. The town-wide prevalence of rust tide (Cochlodinium polykrikoides) may have been a factor in these poor survival and growth results.

The total value of clam production in 2016 was $462,753.

Scallops: In 2015 we tested different gear deployments by overwintering scallops using the usual method (“green blocks”) as well as pearl nets. The blocks proved to be the better option (63% survival vs. 46% in the pearl nets). Overall, of the 117,520 scallops that were overwintered, we seeded 63,911 in the spring (54% survival); less than stellar results but significantly better than the previous year when we had a very harsh winter involving sustained freezing of the overwintering pond.

We began spawning scallops June 1st, with the final spawn on June 7th being the most prolific. We moved approximately 900,000 scallops to the set tanks, but they were kept in the hatchery a bit too long, leading to a significant die-off. In all, we moved 130,000 to the field for growout and overwintered 76,800. After conferring with Mike Patricio at Cornell Cooperative Extension about

1 the die-off, we’ve decided to try their new nursery method in 2017. We will construct smaller- mesh upwellers for use in 2017. This will allow us to move scallops into upwelling earlier (when they’re smaller), with minimal time in set tanks and downwelling, both methods seemingly problematic when it comes to scallop survival.

Our total scallop production value for 2016 was $5,136.

In all, over 50 million shellfish (valued at $759,442) were disseminated in 2016.

Staffing: Barley Dunne, Kate Rossi-Snook, and Pete Topping continued as the full-time Hatchery team, and Jeremy Gould was part-time seasonal Maintenance Mechanic. Adam Younes continued to fill the position of Environmental Aide for the full season. For the summer we had Sam Younes and Tali Friedman as additional Environmental Aides. Carissa Maurin left the Hatchery to pursue graduate school at University of New England. For the first summer since 2011 we were without Shelby Joyce who graduated from University of Miami and began working at Muscongus Bay Aquaculture, ME in May. We wish both Carissa and Shelby the best of luck!

Pete pulling oyster bags for seeding

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2017 Operating Plan

A tote of clams from the field

Target Species: Eastern Oyster (Crassostrea virginica) Hard Clam (Mercenaria mercenaria) Bay Scallop (Argopecten irradians)

Projected Seed/ Oysters: 6-8 million, Spawns: 2/14, 2/28, 3/14 Overwintering Clams: 5-6 million, Spawns: 3/30, 4/13, 4/27 Production: Scallops: 300,000, Spawns: as natural conditioning permits

Permit Status: All East Hampton Shellfish Hatchery marine hatchery and off bottom grow-out permits are in place for the 2017 season

Additional Continue breeding resistant oysters to overcome disease issue Operations/ of 2015 and 2016 Goals: Continue the enumeration of the efficacy of seeding via surveys, especially for clams.

Increase yields/reduce losses in clam growout by trying soft bottom bags.

Expand oyster gardening program

Request fourth full-time staff member

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2016 Oyster Production

Oyster Spawn and Culture Summary

O1 Cohort O2 Cohort O3 Cohort O4 Cohort Dates # Oysters (x106) Dates # Oysters (x106) Dates # Oysters (x106) Dates # Oysters (x106) Spawn 2/22-2/23 13.20 2/25 47.30 3/10 102.90 3/29 102.00 Set Tanks *Early spawn in broodstock 3/7-3/11 9.97 3/25 10.97 4/13 4.36 Downwelling - Hatchery conditioning tank; combined 3/22 5.92 4/5 4.45 4/21 1.69 Set Success salvaged larvae with O2 59% 41% 39% Upwelling - Nursery cohort* 4/12 2.90 4/20 3.10 4/29 1.78 All Cohorts Dates # Oysters (x106) Total to Upwelling 4/12-4/29 7.78 Field Growout 5/26-6/27 5.18 Seeded Culls (<30mm) 6/16-9/9 2.30 Seed (30+mm) 8/30-10/17 1.34 2016 Total Oysters Culled/Seeded: 3.64

Oyster Discards and Culls

Hatchery Discards Nursery and Field Culls Sieve Number Approximate Size Number Sieve Size Number 325 >45um 0 ≤#20 1,260,000 270 >53um 10,860,000 ≤2.0mm 0 230 >63um 51,100,000 ≤2.4mm 500,000 200 >75um 72,300,000 ≤3.4mm 1,196,150 170 >90um 13,100,000 ≤5/16" 1,271,812 140 >106um 26,500,000 <5/8" 194,720 120 >125um 3,500,000 Total Oyster Culls: 4,422,682 100 >150um 18,330,000 80 >180um 1,840,000 70 >212um 4,412,000 60 >250um 0 Total Oyster Hatchery Discards: 201,942,000 Marketable Total: 6,252,000

Islip/Great Atlantic Shellfish Farms was experiencing multiple mass hatchery die-off events, so we donated 136,280,000 of our hatchery discards and 1,260,000 nursery discards to help get their production on track. Stony Brook Southampton received 8,502,000 hatchery discards. We sold 865,540 field culls: 548,540 to Cornell Cooperative Extension, and 317,000 to local oyster farmer Chuck Weimar. The total value of our oyster donations and sales this year was $58,724.

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Oyster Distribution

Nursery Field Sizes (mm) Seed Totals Culls Culls Average Range Accabonac Harbor 0 0 296,724 296,724 49.9 23 - 74 Hog Creek 0 0 72,090 72,090 51.8 35 - 61 Lake Montauk 0 0 256,818 256,818 52.4 31 - 75 Napeague Harbor 0 600,992 300,511 901,503 30.3 8 - 67 Three Mile Harbor 1,696,150 0 334,920 2,031,070 36.6 2 - 64 Northwest Creek 0 0 78,400 78,400 49.9 37 - 63 Donations/Research 1,260,000 0 0 1,260,000 -- 0.5-1.0 Sales 0 865,540 0 865,540 16.3 5 - 37 Totals 2,956,150 1,466,532 1,339,463 5,762,145 Average 2.5 17.4 49.5 Sizes (mm) Range 0.5 - 3.4 5 - 37 28 - 75

Please refer to Appendix I (page 40) for the 2016 Harbor Seeding Maps.

Oyster Overwintering

We did not overwinter oysters in 2016.

Oyster Market Values (includes sales)

Production Market Values Size (mm) Value/1000 Quantity Value Total Value Disca rds >180um $3.01 6,252,000 $18,834.15 $18,834 0.5-1.0 $6.44 1,260,000 $8,111.25 Nursery and 1.1-4.0 $10.75 1,696,150 $18,233.61 $75,693 Field Culls 8.1-12.0 $24.59 539,312 $13,263.70 16.1-25.0 $38.92 927,220 $36,084.31 40.1-45.0 $103.00 467,240 $48,125.72 Seed $185,501 45.1-60.0 $157.50 872,223 $137,375.12 Seeding & 0.5-60.0 $2.00 5,762,145 $11,524.29 $11,524 Handling Total 2016 Oyster Production Market Value: $291,552

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2016 Hard Clam Production

Hard Clam Spawn and Culture Summary

C1 Cohort C2 Cohort C3 Cohort Dates # Clams (x106) Dates # Clams (x106) Dates # Clams (x106) Spawn 4/7 48.80 4/21 12.00 5/5 94.00 Set Tanks 4/22-4/25 5.66 5/6 3.46 5/20 6.38 Transition Tanks - Hatchery 5/2-5/6 11.36 5/20-5/23 6.83 5/31-6/2 6.41 Upwelling - Nursery 5/12-5/19 12.25 5/27-6/6 9.12 6/8-6/16 3.62 All Cohorts Dates # Clams (x106) Total to Upwelling 5/12-6/14 24.99 Seeded Culls (<10mm) 6/9-9/2 31.16 Field Growout 6/23-8/3 6.78 Overwintering 9/28-10/4 0.90 Seed (10+mm) 8/1-10/4 3.74 2016 Total Clams Culled/Seeded/Overwintered: 35.80

Hard Clam Discards and Culls

Hatchery Discards Nursery and Field Culls Sieve Number Approximate Size Number Sieve Size Number 230 >63um 0 ≤#20 23,853,750 200 >75um 38,200,000 ≤1.0mm 4,458,750 170 >90um 35,380,000 ≤2.0mm 800,000 140 >106um 12,500,000 ≤2.4mm 70,000 120 >125um 16,700,000 ≤3/16" 1,981,440 100 >150um 0 Total Clam Culls: 31,163,940 80 >180um 374,000 70 >212um 185,000 60 >250um 1,669,000 50 >300um 5,664,500 40 >425um 3,619,000 Total Clam Hatchery Discards: 114,291,500 Marketable Total: 11,511,500

We did not donate any hatchery discards this year. Instead, over 11.5 million viable juvenile clam discards, sizes #80 – #40, were seeded to just south of the Nursery and east of the Lazy Point boat ramp. Stony Brook Southampton received 1,000 field culled clams.

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Hard Clam Distribution

Nursery Field from 2015 Sizes (mm) Seed Totals Culls Culls Cohort Average Range Accabonac Harbor 0 776,064 451,776 121,072 1,348,912 11.2 4 - 19 Hog Creek 0 0 0 50,400 50,400 13.8 -- Lake Montauk 3,540,000 0 1,026,000 0 4,566,000 6.4 <1 - 16 Napeague Harbor 6,200,000 428,312 882,726 121,072 7,632,110 10.0 <1 - 19 Three Mile Harbor 19,442,500 776,064 869,401 157,920 21,245,885 6.9 <1 - 15 Northwest Creek 0 0 513,704 0 513,704 11.7 7 - 16 Overwintering 0 0 895,800 0 895,800 12.3 7 - 16 Donations/Research 0 1,000 0 0 1,000 7.3 4 - 10 Totals 29,182,500 1,981,440 4,639,407 450,464 36,253,811 Average -- 7.3 11.7 13.2 Sizes (mm) Range <1 - <2.4 4 - 10 6 - 19 --

Please refer to Appendix I (page 40) for the 2016 Harbor Seeding Maps.

Hard Clam Overwintering

Overwintering Stocking and Retrieval 2016 Overwintering Stocking Northwest Creek: Stocked 10/27/15 1,034,830 Northwest Creek: Total Stocked 9/28/16 850,800 average size (mm) 13.0 5/16" clams 247,360 Seeded 9/12/16-9/13/16 450,464 average size (mm) 13.5 average size (mm) 13.2 3/16"x3 clams 603,440 Percent survival 44% average size (mm) 10.8 Percent growth 1% Test Sites: Total Stocked 10/4/16 45,000 average size (mm) 12.5 Accabonac Harbor 15,000 Napeague Harbor: Pond Intet 15,000 Napeague Harbor: Pond 15,000

The 2015 overwintered clams experienced low survival (44%) and very poor growth rate (1%). In response, we overwintered some clams in three additional test sites this year; one in Accabonac Harbor, and two in Napeague Harbor (inside Pond of Pines and in the channel leading into the pond). If survival and/or growth is better in any of those sites than Northwest Creek, we may change our clam overwintering location. However, as mentioned above, poor survival and growth may have been due to the presence of Cochlodinium sp. across the region.

In total, the 2016 overwintered clams were stocked in 368 bags at ~2,500 clams per bag. In an effort to reduce losses to washout & predation we replaced the old style boxed end bags with sealed end bags.

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Hard Clam Market Values

Production Market Values Size (mm) Value/1000 Quantity Value Total Value Disca rds >180um $2.00 11,511,500 $23,023.00 $23,023 0.5-1.0 $5.75 28,312,500 $162,796.88 1.1-2.0 $8.63 800,000 $6,900.00 Culls $215,755 2.1-3.0 $14.00 70,000 $980.00 6.1-8.0 $22.75 1,981,440 $45,077.76 Seed 10.1-16.0 $36.32 3,743,607 $135,973.15 $135,973 2 Year Old Seed 12.1-16.0 $38.38 450,464 $17,286.56 $17,287 (2015 Cohort) Seeding & 0.5-25.0 $2.00 35,358,011 $70,716.02 $70,716 Handling Total 2016 Clam Production Market Value: $462,753

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2016 Bay Scallop Production

Bay Scallop Spawn and Culture Summary

S1 Cohort S2 Cohort S3 Cohort Dates # Scallops (x106) Dates # Scallops (x106) Dates # Scallops (x106) Spawn 6/1 15.12 6/2 3.60 6/7 21.3 All Cohorts Dates # Scallops (x106) Set Tanks 6/13-6/17 0.90 Downwelling - Nursery 6/20 0.75 Upwelling - Nursery 6/24-7/1 0.36 Field Growout 7/26-8/1 0.13 Seeded Culls (<20mm) 8/1 0.04 Overwintering 9/26-9/27 0.08 2016 Total Scallops Culled/Seeded/Overwintered: 0.12

Bay Scallop Discards and Culls

We kept our hatchery discards to a minimum this year. We seeded 38,250 culls off the Nursery dock, and donated 3,879 field culls to Stony Brook Southampton.

Hatchery Discards Nursery and Field Culls Sieve Number Approximate Size Number Sieve Size Number 325 >45um 0 ≤2.4mm 38,250 270 >53um 730,000 <20mm 3,879 230 >63um 1,600,000 Total Scallop Culls: 42,129 200 >75um 800,000 170 >90um 1,000,000 140 >106um 0 120 >125um 0 100 >150um 0 80 >180um 0 70 >212um 0 60 >250um 0 50 >300um 0 40 >425um 0 Total Scallop Hatchery Discards: 4,130,000 Marketable Total: 0

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Bay Scallop Distribution

Nursery Field from 2015 Sizes (mm) Seed Totals Culls Culls Cohort Average Range Accabonac Harbor 0 0 0 0 0 -- -- Hog Creek 0 0 0 0 0 -- -- Lake Montauk 0 0 0 0 0 -- -- Napeague Harbor 0 0 0 32,909 32,909 35.8 30 - 45 Three Mile Harbor 38,250 0 0 31,002 69,252 19.0 <2.4 - 45 Northwest Creek 0 0 0 0 0 -- -- Pond of Pines Overwintering 0 0 76,800 0 76,800 -- -- Donations/Research 0 3,879 0 0 3,879 11.8 8 - 16 Totals 38,250 3,879 76,800 63,911 182,840 Average <2.4 11.8 -- 35.8 Sizes (mm) Range -- 8 - 16 -- 30 - 45

Please refer to Appendix I (page 40) for the 2016 Harbor Seeding Maps.

Bay Scallop Overwintering

Overwintering Stocking and Retrieval 2016 Overwintering Stocking Pond of Pines Blocks: Stocked 10/19/15 57,920 Pond of Pines: Stocked 9/26/16-9/27/16 76,800 average size (mm) 29.8 average size (mm) --- Seeded 5/10/16-5/24/16 36,622 percent survival 63% Pond of Pines Pearl Nets: Stocked 10/20/15 59,600 average size (mm) 28.6 Seeded 5/10/16-5/24/16 27,289 percent survival 46% Total Stocked 117,520 average size (mm) 29.2 Total Seeded 63,911 average size (mm) 35.8 Total percent survival 54% Total percent growth 22%

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Bay Scallop Market Values

Production Market Values Size (mm) Value/1000 Quantity Value Total Value Disca rds >180um $11.00 0 $0.00 $0 1.5-2.4 $16.50 38,250 $631.13 Culls 6.1-10.0 $32.00 975 $31.20 $770 10.1-20.0 $37.00 2,904 $107.45 Seed 20.1-45.0 $58.33 0 $0.00 $0 2 Year Old Seed 26.0-45.0 $65.00 63,911 $4,154.22 $4,154 (2015 Cohort) Seeding & 1.5-45.0 $2.00 106,040 $212.08 $212 Handling Total 2016 Scallop Production Market Value: $5,136

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2016 Projects and Cooperative Research

East Hampton Shellfish Education and Enhancement Directive (EHSEED)

Co-sponsors East Hampton Town Shellfish Hatchery South Fork Natural History Museum Veterinarians International

Summary A corps of eco-conscious South Fork residents collaborated with the East Hampton Town Shellfish Hatchery to expand shellfish education and enhancement programs in East Hampton town waters by forming an oyster-growing community cooperative, or oyster growing program. EHSEED creates opportunities for local citizens to learn about and help the threatened ecosystem that is part of the vital Peconic Estuary system, resulting in educated and dedicated stewards of the environment. EHSEED is financed by a combination of individual and family memberships in the project. The Shellfish Hatchery staff implement the educational program and provide the training. Lectures and workshop offerings take place at the Shellfish Hatchery’s facilities in Montauk and Springs. The South Fork Natural History Museum (SoFo) enhances awareness and membership in the program by disseminating information to its members. Veterinarians International, a non- profit organization committed to enhancing the health of humans, animals and the environment, promotes the project to its affiliates and local residents. To participate in EHSEED, each member/family must be Town of East Hampton residents and pay a membership fee of $250. This fee entitles them to training, 1,000 oyster seed (one half will be seeded to Town waters in the spring, under the direction of the Hatchery, the other is for them to keep), and the gear required to grow the oysters to harvest size (which should occur by the end of the second growing season).

The 2016 pilot program hosted 15 members. The season kicked off with a fundraiser/meet-and- greet at Bay Kitchen Bar on Three Mile Harbor, and was followed by a series of four lectures: • Shellfish biology and broodstock conditioning • Larvae and algae culture • Field culture with seed distribution to members • Shellfish disease and proper handling Five gear maintenance and size grading workshops were held throughout the season, with a final meeting for overwintering and seeding. Participants will return in spring to continue growing and caring for their oysters until harvest size, at which point they can harvest them or continue to grow them. EHSEED hopes to expand to include an additional 15 members (30 members total) for the 2017 season.

The EHSEED site at the Nursery EHSEED oyster gardeners hard at work

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Hatchery Assisted Student Research Projects:

Battle of the Bivalves

Participant Anni Spacek, Montauk Elementary School 8th grade

Abstract The purpose of the experiment was to discover what the best filterer is of Fort Pond baywater. Atlantic Oysters, Quahog Clams, or shells? Using the East Hampton Town Shellfish Hatchery, there were six tanks; two with 12 oysters each, two with 12 clams each and two with 610.10 grams of shells each. Every week each tank was tested for pH, ammonia, nitrite and nitrate. Every three days instant marine algae paste was added to the water in all tanks. This process continued for ten weeks, from October 18th to January 1st. The conclusion that was reached is that the clams filtered nitrite and nitrate the best, and had the most beneficial impact on Fort Pond Bay water. Overall, the tanks containing the clams had the lowest amounts of contaminants.

Results: 1st prize in Montauk School Science Fair, qualifies for entry into the L.I. Science Congress Regional Fair in April

East Hampton Shellfish Hatchery’s role: provided shellfish, mentoring, equipment and space to conduct work

Oyster Murder Mystery

Participant Sophia Botero, Montauk Elementary School 8th grade

Abstract Fishing for sea creatures is big in our community here in Montauk. It is a great factor in our economy and business. I studied oysters to see how people’s habits affect them. Pollutants can affect the sea animals and their habitat. I’m doing this experiment because I want to raise awareness to how we treat our sea animals, and how it affects both us and them. If oysters exposed to runoff pollutants is related to oyster mortality or illness, then oyster beds adjacent to roads, homes or golf courses will have high mortality rates. I found that if I expose oysters to small amounts of pollutants, they die over time. It has been proven that oysters filter out most of the nitrogen that is in the water which is a good thing. If we eliminate the oysters then the nitrogen level will increase. Research is improving our understanding of how excess nitrogen affects air, water and soil quality, puts pressure on ecosystems and biodiversity, leads to human health risks, and affects the global climate. Because of human modification of the nitrogen cycle, thresholds for reactive nitrogen have already been exceeded in some places – with the risk of bringing about abrupt or irreversible environmental changes. Organisms require oxygen to grow and reproduce.

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‘Dead zones’ are oxygen-depleted (hypoxic) areas that result from over-enrichment of waters with nutrients (especially nitrogen and phosphorus), for example from fertilizer run-off, industrial waste and sewage. Reported cases of coastal dead zones have doubled in each of the last four decades.

Results: 2nd prize in Montauk School Science Fair, qualifies for entry into the L.I. Science Congress Regional Fair in April

East Hampton Shellfish Hatchery’s role: provided shellfish, mentoring, equipment and space to conduct work

Hatchery Assisted External Professional Research Projects:

Understanding the effects of predicted ocean conditions as a result of climate change on juvenile shellfish

Participant Alexandra Stevens, Marine Sciences Program, Stony Brook Southampton

Summary Two experiments were conducted focusing on future climate change risks on shellfish generously supplied by East Hampton Town Shellfish Hatchery. Each of the two experiments used different shellfish species (~500 of each), including hard clams and bay scallops, all of which were obtained during their juvenile stage. These experiments used juvenile shellfish because it is one of the more vulnerable stages for these economically important shellfish. The shellfish were grown in conditions similar to the predicted climate change and warming in our local bays and oceans. Both experiments encompassed an ambient condition, a low pH condition, a low dissolved oxygen (D.O.) condition, and combined low pH and low dissolved oxygen conditions. The ambient condition represented current ocean/bay conditions (pH: ~7.9-8 & DO: ~6mg/L). Low pH conditions represented future predictions for increasing acidification in our local bays and oceans (pH: ~7.2 & DO: ~6mg/L). Low dissolved oxygen represented future predictions on less dissolved oxygen/increasing hypoxia in our local bays and oceans (pH: ~7.9-8 & DO: ~2mg/L). The fourth condition represented both ocean acidification and hypoxia in future waters (pH: ~7.2 & DO: ~2mg/L). These experiments also placed shellfish under varied temperature conditions. Typically at an ambient temperature of 24˚C as well as a higher temperature (~30-31˚C) to represent future predictions of increasing ocean temperatures. The shellfish were then placed in buckets (4 replicates of each) for each condition (four conditions) over a period of time to measure growth rate and survival throughout the experiment. Daily feeding from our lab's algae and measurements of chemistry were taken. Once the data is analyzed the goal of these experiments will be to better understand how juvenile shellfish survive and grow under future ocean conditions.

On a much smaller scale, two brown tide (BT) experiments were conducted, using recently settled juvenile clams and scallops from East Hampton Town Shellfish Hatchery. These experiments were conducted during early summer BT blooms in our local bays. Water was collected at locations where BT was abundant and used in the experiment. The experimental design used BT water and filtered water (i.e. control) to compare the growth and survival of these local bivalve species under harmful conditions. This experiment also included the macroalgae Ulva, to study the possible mitigation effect Ulva may have on brown tide. The juvenile clams and scallops were placed in

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the following treatments for a few weeks (however long the bloom lasted): control, brown tide, brown tide and Ulva. All conditions had four replicates, and were fed daily from our lab's algae.

Results: in progress

East Hampton Shellfish Hatchery’s role: provided juvenile shellfish

Shinnecock Bay Restoration Project oyster reef construction

Participant Andrew W. Griffith, Marine Sciences Program, Stony Brook Southampton

Summary The East Hampton Town Shellfish Hatchery provided ~ 3 million oyster larvae for the Shinnecock Bay Restoration Project (www.shinnecockbay.org). Prior to metamorphosis, larvae were transferred to the Stony Brook Southampton Marine Sciences center and carefully added to large volume aquaria (> 1000 L, n = 3) containing Vexar mesh bags filled with surf clam (Spisula solidissima) shell. Oyster larvae were allowed to set directly onto shell within mesh bags. These shell bags containing live oysters will be used to construct oyster reefs in the western regions of Shinnecock Bay. Constructed reefs will provide substrate for future oyster recruitment as well as establish source populations for enhanced recruitment in nearby areas. Established oyster reefs will provide critical habitat for ecologically and economically significant species of marine organisms as well as improve water quality through the removal of particulate organic matter and the sequestration of excess nutrients from the water column.

Results: in progress

East Hampton Shellfish Hatchery’s role: provided oyster larvae

Impacts of climate change and ocean acidification on economically important shellfish in New York: are there effective mitigation and adaptation measures?

(Adapted from NYSG Progress Report R/FBM-38)

Participants Christopher J. Gobler Andrew Griffith Ryan B. Wallace

Objective 2: Document the co-occurrence of acidification, low oxygen, and/or thermal stress in NY shellfish hatcheries, aquaculture regions, and estuaries.

Field Sampling Beginning in early June of 2016, horizontal surveys were conducted using small craft within the South Shore Estuary Reserve (SSER) from Shinnecock Bay through Great South Bay on a monthly basis. Additional cruises were conducted in Peconic Bay aboard the R/V Paumanok on a monthly basis. Additionally, discrete sampling coupled with sensor arrays (YSI EXO2) were deployed at

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two shellfish hatcheries, Town of East Hampton and Town of Islip, in the spring. For comparative purposes, sampling and sensors were also placed in the Southampton Marine Lab during this same period. Finally, discrete sampling methods and sensors (YSI EXO2 and SesaFET) were placed at multiple locations throughout both the Peconic Estuary and Shinnecock Bay. These arrays were employed to characterize the seasonal and diurnal variation of dissolved oxygen, DIC and pH in distinct coastal habitats during spring, summer, and fall months.

Results The deployment of sensors at multiple shellfish hatcheries gave us valuable baseline data to build upon this coming field season with the expectation of additional hatcheries becoming involved in the monitoring program this coming season. The monitoring of Southampton Marine Station incurrent water proved to be valuable for comparison as we were able to acquire consistent diurnal patterns (Fig.7) April - May, 2016. The pH during this time period ranged from 7.8 - 8.3, while the DO ranged from 7 – 14 mg L-1 (Fig. 7). Sensors at the East Hampton and Islip hatcheries were installed in the upwelling chambers in which water is circulated shortly after the shellfish have attached. In the East Hampton hatchery incurrent water is from Block Island Sound and the pH range was 7.7 – 8.2 (Fig. 7). This water was temperature controlled at 25ºC and significantly warmer than ambient water (~10 ºC). DO remained relatively stable during this period (Fig 7; ~7 mg L-1). The Islip Hatchery acquires their water from a saltwater well and the water is then stored in a holding tank before undergoing an iron filtration process. The DO at the Islip hatchery was slightly lower and not as stable as the East Hampton hatchery. DO ranged from 5 – 7 mg L-1 and pH ranged from 7.4 – 8.1 (Fig. 7). Even with healthy DO concentrations, pH is significantly depressed at the Islip hatchery during early life stage development when these bivalves are most sensitive to acidification (Barton et al., 2012; Gobler and Talmage, 2013; Waldbusser et al., 2013). Objective 4: Assess the individual and combined effects of the thermal stress, low oxygen, and acidification on the early life stages of NY’s most valuable resource bivalves.

Experimental Methods The experimental approach was designed to expose early life stage bivalves to normal conditions, low pH, high temperature, and low dissolved oxygen conditions in a full factorial design (individually and in all combinations) achieved via the delivery of varying mixtures of CO2 gas, N2 gas and air. Experiments were performed with larval stage and recently settled juvenile stages of NY’s most valuable bivalve fisheries: hard clams (Mercenaria mercenaria), Eastern oysters (Crassostrea virginica), and bay scallops (Argopecten irradians).

Gasses were mixed as outlined in Gobler et al (2014) with gas proportioners used to deliver mixes of CO2 gas, N2 gas, and air to seawater to maintain conditions that are oxic with a normal pH (air only), hypoxic with a normal pH (tanked N2 gas pre-mixed with ~390 µatm CO2 gas), oxic but acidified (tanked 5% CO2 gas and air), and hypoxic and acidified (tanked N2 gas, tanked 5% CO2 gas, and air) gas. Prior studies using these gases have demonstrated that we are able to create conditions which mimic those found in estuaries during summer with regard to oxygen, pH, and pCO2 levels for more than one month (DePasquale et al., 2015; Gobler et al., 2014) as well as temperatures that cover the normal to high range found in estuaries. Temperatures were maintained by placing experimental vessels (10-L polyethylene buckets) in seatables with circulating water maintained via Delta® Star heater-chiller-pump units. For this study, we

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contrasted ideal and stressful conditions for each given parameter for the purposes of identifying organisms that are vulnerable and resistant to extreme levels of each individual parameter and combinations of conditions. Our experimental conditions were (T = 24°C, pH = 8.0, DO = 8 mg L-1) acidified (pH = 7.4), hypoxic (DO = 2 mg L-1), and warm (30°C) in a full factorial design (individually and in all combinations), creating conditions that are present, in shallow, eutrophic NY estuaries today (Wallace et al., 2014) and that will become more common and wide-spread under future climate change conditions (Doney et al., 2012). Experiments were performed for each species (n=3) and stage (n=2, larvae and juvenile; details below) for a total of six experiments by administering conditions of constant temperature, DO, and pH conditions as described above. These experiments (n=6) were repeated (leading to 12 total experiments) by administering levels of temperature, DO, and pH that fluctuate on a diel basis from the control condition to the extreme condition, mimicking changes observed in some shallow, NY estuaries (Fig. 11). Diurnal changes in conditions were produced by using ITT Alcon solenoid valves attached to compressed air tanks and ambient air lines and were controlled with programmable, electronic timers that deliver ambient air by day and mixtures of N2, CO2, and air at night to produce the desired low pH and DO concentrations. Concurrently, heater-chillers were programed to vary temperatures in unison with changes in water chemistry. These devices have generated diurnal changes in water chemistry highly similar to changes observed in shallow, estuarine ecosystems (Fig 12, compare to Fig 3). Actual measurements of chemical conditions present during all experiments were made via the use of data logging dissolved oxygen (Onset), temperature (Onset), and pH probes (Orion) which record the precise levels of DO, pH, and temperature in experimental vessels every 10 minutes.

Both larval and juvenile shellfish were obtained from the East Hampton Town Shellfish Hatchery located in Montauk, NY, which uses broodstock collected from the normoxic, normocapnic regions of the eastern Peconic Estuary. For experiments, each 10-L polyethylene vessel was stocked with either 10,000 D-stage larvae that are less than 24 hours old or 15 juveniles that are less than one month old and thus vulnerable to environmental stressors (Gobler et al., 2014; Green et al., 2009). Larvae were fed a diet of 4 x 104 cells ml-1 of Isochrysis galbana (Tahitian strain: T- Iso) daily, while juveniles will be fed a mix of T-Iso and Tetraselmis chuii. Bivalve food was added to 0.2 µm filtered seawater from Old Fort Pond in Shinnecock Bay, NY, treated with a 1% concentration of an antibiotic solution (Sigma-Aldrich No. 4083, penicillin, streptomycin, neomycin) to deter bacterial growth (Talmage and Gobler, 2009). Water changes were performed twice weekly when, for larvae, contents of the experimental vessels were poured over a 64 µm sieve. Larvae collected on the sieve were transferred to a 50 ml container from which 2 ml was removed and preserved with a 3% solution of formalin. The remaining larvae were returned to experimental vessels while preserved larvae were used to assess mortality, growth, and development at each time point using a dissecting microscope with Nikon DigiSight Color Digital Camera System (DSVi1) and ImageJ software. The length (distance from tip of the umbo to the furthest leading ventral side) of each larva were measured and their developmental stage noted (veliger/pediveliger or metamorphosed). Juvenile bivalve mortality was noted during each biweekly water change and lengths were measured with digital calipers to calculate growth rates. Larval experiments persisted until all individuals have metamorphosed within control vessels whereas juvenile experiments persist for ~ one month spanning the critical period for their growth and survival (Green et al., 2009). The effects of acidification, hypoxia, and thermal stress on the growth and survival of early life stage bivalves were assessed via a three-way analysis of variance (ANOVA) where pH, dissolved oxygen, and temperature were the main effects.

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Results Survival of larval A. irradians was significantly reduced by acidification (Fig. 13). Larvae exposed to the control and intermediate levels of pH (pH = 7.94 ± 0.06 and 7.60 ± 0.03, respectively) experienced 36 ± 2% and 21 ± 11% survival (± standard deviation), respectively, whereas the percentage of larvae that survived in the low pH treatment (pH = 7.33 ± 0.07; survival = 7 ± 5%) was significantly lower than the intermediate (p = 0.026) and control (p = 0.003), but not different than the diurnal treatment (mean pH = 7.47 ± 0.43; survival = 15 ± 3%). Low pH conditions slowed larval growth (Oneway ANOVA; p < 0.001) and delayed metamorphosis (One-way ANOVA; p < 0.001) relative to the control treatment, but there was no effect of intermediate and diurnal pH treatments on growth or development. Larvae grew at a rate of 11 ± 5 μm d-1 in the low pH treatment, 26 ± 4 μm d-1 in the control treatment, 20 ± 2 μm d-1 in the intermediate pH treatment, and 22 ± 3 μm d-1 in the diurnal pH treatment (Fig. 13B). Twelve days postfertilization, 88 ± 5%, 58 ± 22%, and 61 ± 19% of A. irradians larvae had metamorphosed in the control, intermediate, and diurnal treatments while only 13 ± 19 % had metamorphosed in the low treatment (Fig. 13C).

Low pH (pH = 7.29 ± 0.06) reduced survival (One-way ANOVA; p = 0.015) of larval M. mercenaria though there was no effect of intermediate (pH = 7.58 ± 0.04) or diurnally fluctuating pH (mean pH = 7.54 ± 0.36). Larvae reared under control (pH = 7.91 ± 0.02) , intermediate, and diurnal pH conditions had survival rates of 27 ± 3%, 23 ± 2%, and 25 ± 5% respectively whereas survival of larvae in the low pH treatment was 15 ± 5%, significantly lower than all other treatments (Fig. 14A). Growth and development of M. mercenaria larvae did not differ significantly among experimental treatments (Fig. 14B, C).

Survival was significantly reduced in juvenile A. irradians exposed to low pH (pH = 7.13 ± 0.03; One-way ANOVA; p = 0.016), intermediate pH (pH = 7.49 ± 0.04; p = 0.003), and 12 diurnally fluctuating pH conditions (mean pH = 7.57 ± 0.43; p = 0.001). In the control treatment (pH = 7.92 ± 0.05), 92 ± 6 % of individuals survived whereas low, intermediate, and diurnally fluctuating pH treatments experienced 57 ± 25%, 47 ± 9%, and 40 ± 12% survival, respectively, but were not significantly different from each other (Fig. 15A). There were no pH effects on the growth rates of juvenile A. irradians (Fig. 15B). The differing levels of pH used in experiments did not significantly alter the survival and growth of juvenile M. mercenaria (Fig. 16A, B).

Diurnal exposure to low pH and low DO (mean pH = 7.61 ± 0.26; mean DO = 4.11 ± 2.80 mg L- 1) significantly reduced survival of larval A. irradians (One-way ANOVA; p = 0.023), but chronically low and intermediate pH and DO conditions (pH = 7.22 ± 0.05; DO = 1.38 ± 0.45 mg L-1 and pH = 7.48 ± 0.05; DO = 4.08 ± 0.41 mg L-1, respectively) had no effect. The percent survival of larval A. irradians in the control (pH = 7.89 ± 0.00; DO = 6.87 ± 0.25 mg L-1), intermediate, and low pH and DO treatments was 15 ± 10%, 17 ± 6%, and 6 ± 4%, respectively, while survival in the diurnally fluctuating treatment was 3 ± 1% (Fig. 17A). Both the low and diurnal treatment slowed growth (One-way ANOVA; p < 0.001) from 13 ± 1 μm d-1 and 13 ± 1 μm d-1 in the control and intermediate pH-DO treatments to 7 ± 1 μm d-1 and 10 ± 0.4 μm d-1 in the low and diurnal pH-DO treatments (Fig. 17B). Continuously low pH and DO also significantly delayed development (One-way ANOVA; p = 0.016), while exposure to intermediate and diurnally fluctuating pH and DO did not. After 15 days, 32 ± 8%, 36 ± 4%, and 32 ± 13% of larvae had metamorphosed in the control, intermediate, and diurnal pH-DO treatments, whereas only 13 ± 3% had metamorphosed in the low pH-DO treatment (Fig. 17C).

Exposure of larval M. mercenaria to low (pH = 7.24 ± 0.04; DO = 1.32 ± 0.30 mg L-1) and diurnally fluctuating pH-DO (mean pH = 7.41 ± 0.34; mean DO = 4.02 ± 3.00 mg L-1) significantly reduced

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their survival to 7 ± 2% and 5 ± 2% compared to the control (pH = 7.87 ± 0.03; DO = 6.92 ± 0.13 mg L-1) and intermediate (pH = 7.43 ± 0.03; DO = 3.92 ± 0.34 mg L-1) pH survival which was 13 ± 2% and 8 ± 2%, respectively (One-way ANOVA; p = 0.032; p = 0.001; Fig. 18A). Continuously low pH and DO conditions slowed the growth of M. mercenaria larvae to 9 ± 2 μm d-1 compared to 13 ± 0.4 μm d-1, 12 ± 1 μm d-1, and 12 ± 2 μm d-1 in the control, intermediate (pH = 7.43 ± 0.03; DO = 3.92 ± 0.34 mg L-1), and diurnally fluctuating pH-DO treatments (One-way ANOVA; p < 0.05; Fig. 18B). Delays in development were observed when M. mercenaria larvae were exposed to lowered pH and DO (One-way ANOVA; p < 0.05). Eleven days post-fertilization, 61 ± 4% of larvae had metamorphosed in the control treatment, whereas only 41 ± 7%, 28 ± 9%, and 24 ± 7% had metamorphosed in the intermediate (p = 0.019), diurnal (p < 0.001), and low pH-DO (p < 0.001) treatments respectively (Fig. 18C).

There was a significant negative effect of pH (Two-Way ANOVA; p < 0.001), and DO (p < 0.001) on survival of A. irradians larvae and an antagonistic interaction between both factors (p < 0.05) with all manipulated conditions significantly reducing survival relative to the control condition. Percent survival for the control, low pH, low DO, low pH-DO, diurnal pH, diurnal DO, and diurnal pH-DO conditions was 38 ± 2%, 15 ± 4%, 25 ± 6%, 5 ± 6%, 12 ± 3%, 17 ± 5%, and 7 ± 2% (Fig. 19A). Survival under continuously low DO (pH = 7.91 ± 0.02; DO = 2.33 ± 0.64 mg L-1) was higher than both the continuously low pH (pH = 7.20 ± 0.09; DO = 6.96 ± 0.74 mg L-1; p = 0.006) and diurnal pH-DO (mean pH = 7.58 ± 0.25; mean 14 DO = 4.28 ± 2.90 mg L-1; p < 0.001) conditions, though still significantly lower than the control treatment (pH = 7.91 ± 0.02; DO = 6.87 ± 0.33 mg L-1; p = 0.009). The antagonistic interaction between DO and pH was most apparent in the diurnal pH-DO treatment where the survival (7 ± 2%) was higher than would have been predicted by the reductions in survival in the diurnal pH and diurnal DO treatments separately (12 ± 3% and 17 ± 5%, respectively).

Growth rates of A. irradians larvae were affected by both pH (Two-way ANOVA; p < 0.001) and DO (p < 0.001). There was no interaction between the factors. Larvae experienced significantly slowed growth under all manipulated conditions, except for the diurnally fluctuating pH treatment (mean pH = 7.47 ± 0.23; DO = 6.80 ± 0.57 mg L-1; growth rate = 10 ± 2 μm d-1; Fig. 19B). Control larvae grew at a rate of 13 ± 1 μm d-1 while rates were slowed to 6 ± 1 μm d-1 in low pH-DO (p < 0.001), 7 ± 1 μm d-1 in low pH (p < 0.001), 8 ± 1 μm d-1 in diurnal pH-DO (p < 0.001), 9 ± 1 μm d-1 in diurnal DO (p = 0.018), and 9 ± 2 μm d-1 in low DO (p = 0.024). Both pH (p < 0.001) and DO (p < 0.001) affected development of A. irradians larvae (Two-way ANOVA). Fourteen days postfertilization, 67 ± 5% of larvae had metamorphosed under control conditions (Fig. 19C). Continuously low pH (p < 0.001) and continuously low DO (p = 0.026) reduced metamorphosis to 35 ± 9% and 48 ± 10% respectively, but diurnal exposure of low pH and of low DO did not alter the fraction of larvae that had metamorphosed. Metamorphosis was delayed in the low pH-DO treatment (p < 0.001) to 14 ± 6% and to a significantly lesser extent in the diurnal pH-DO treatment (38 ± 9%; p < 0.001).

Survival of larval M. mercenaria was significantly reduced by pH (Two-way ANOVA; p < 0.001) but not DO, and there was an antagonistic interactive effect of these two factors (p < 0.05). Under control (pH = 7.97 ± 0.07; DO = 7.13 ± 0.20 mg L-1), chronically low DO (pH = 7.92 ± 0.08; DO = 2.64 ± 0.40 mg L-1), and fluctuating low DO conditions (pH = 7.92 ± 0.06; mean DO = 4.44 ± 2.70 mg L-1), 28 ± 2%, 24 ± 3%, and 27 ± 5% of larvae survived to the end of the experiment, while survival was significantly reduced to 1 ± 0.4%, 2 ± 1%, 1 ± 1%, and 5 ± 3% in the low pH (pH = 7.21 ± 0.10; DO = 7.14 ± 0.22 mg L-1; p < 0.001), diurnal pH (mean pH = 7.43 ± 0.65; DO = 7.47 ± 0.24 mg L-1; p < 0.001), low pH-DO (pH = 7.22 ± 0.10; DO = 1.90 ± 0.38 mg L-1; p <

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0.001), and diurnal pH-DO (mean pH = 7.55 ± 0.56; mean DO = 4.49 ± 2.60 mg L-1; p < 0.001) treatments (Fig. 20A). There was antagonism between pH and DO as the survival in the combined diurnal treatment was higher than would have been predicted by either individual treatment for both the static and diurnal treatments. There was an effect of pH (Two-way ANOVA; p < 0.05) and DO (p < 0.05) on the growth of M. mercenaria larvae, with the growth rates in the control, low pH, low DO, low pH-DO, diurnal pH, diurnal DO, and diurnal pH-DO treatments being 9 ± 0.4 μm d-1, 9 ± 1 μm d-1, 8 ± 1 μm d-1, 8 ± 0.4 μm d-1, 8 ± 1 μm d-1, 9 ± 0.4 μm d-1, and 8 ± 1 μm d-1 although post-hoc multiple comparisons did not identify differences among treatments (Fig. 20B). Development of M. mercenaria larvae was affected by pH (Two-way ANOVA; p < 0.001), DO (p < 0.001), and there was an antagonistic interaction between these two factors (p < 0.001). Fewer larvae metamorphosed 17 days post- fertilization in all pH and DO treatments compared to 80 ± 6% that had metamorphosed in control conditions (p < 0.001; Fig. 20C). There was an antagonistic effect of pH and DO on development with 21 ± 6% and 31 ± 7% of larvae reaching metamorphosis in the chronically low pH (p < 0.001) and chronically low DO (p < 0.001) treatments and 7 ± 2% in the chronically low pH-DO (p < 0.001) treatment, a value higher than would have been predicted by the individual treatments. Although fewer larvae developed to metamorphosis in the low DO and diurnal DO (25 ± 5%; p < 0.001) treatments than the control, there were significantly more metamorphosed larvae in these two treatments than the diurnal pH (9 ± 4%; p < 0.001; p = 0.002), low pH-DO (p < 0.001; p < 0.001), and diurnal pH-DO (11 ± 3%; p = 0.001; p = 0.034) treatments.

DO and pH significantly reduced the survival of C. virginica larvae (Two-way ANOVA; p < 0.05 and p < 0.001, respectively) and there was no interaction between these factors. Survival was reduced from 15 ± 5% under control conditions (pH = 7.85 ± 0.04; DO = 7.04 ± 0.16 mg L-1), to 3 ± 1%, 5 ± 4%, and 5 ± 2% in low pH (pH = 7.16 ± 0.07; DO = 6.98 ± 0.16 mg L-1; p < 0.001), low pH-DO (pH = 7.18 ± 0.07; DO = 1.87 ± 0.41 mg L-1; p = 0.015), and diurnal pH-DO (mean pH = 7.50 ± 0.23; mean DO = 4.36 ± 2.80 mg L-1; p = 0.017) treatments (Fig. 21A). There was, however, no significant differences between control conditions and diurnal fluctuations in pH (mean pH = 7.54 ± 0.26; mean DO = 7.54 ± 0.18 mg L-1; survival = 6 ± 1%) as well as between diurnal and continuous low DO on survival percentages (mean pH = 7.99 ± 0.09; mean DO = 5.14 ± 2.10 mg L-1; survival = 14 ± 5%; pH = 7.83 ± 0.05; DO = 2.50 ± 0.71 mg L-1; survival = 21 ± 7%, respectively); these survival rates were all significantly higher than the low pH, low pH-DO, and diurnal pH-DO treatments. Finally, the percent survival of C. virginica larvae was significantly higher in the low DO treatment than the diurnal pH treatment (p = 0.002). There was an overall effect of pH (Twoway ANOVA; p < 0.001), DO (p < 0.05), and an antagonistic interactive effect of pH and DO (p < 0.05) on the growth rates of C. virginica larvae. Growth rates were reduced from 1 ± 0.1 μm d-1 in control conditions to 0.7 ± 0.1 μm d-1, 0.3 ± 0.2 μm d-1, 0.2 ± 0.1 μm d-1, and 0.4 ± 0.4 μm d-1 in low pH (p = 0.014), diurnal pH (p < 0.001), low pH-DO (p < 0.001), and diurnal pH-DO (p < 0.001) conditions (Fig. 21B). Growth rates of C. virginica larvae exposed to chronically low or diurnal fluctuations of DO (1 ± 0.4 and 1 ± 0.2 μm d-1, respectively) did not differ from the control treatment. The antagonistic effect of pH and DO on C. virginica growth rates was most obvious in the diurnal treatments where exposure to diurnally low pH and DO yielded growth rates higher than would have been predicted by the individual treatments. Metamorphic state was not quantified for C. virginica larvae.

Interactions We collaborated with both the Town of East Hampton and Town of Islip shellfish hatchery. Martin Byrnes, Environmental Coordinator at the Town of Islip Hatchery allowed us to monitor water quality within the facility and John Dunne, Director at the East Hampton Hatchery also allowed

20 us to collect water quality data. Additionally, East Hampton Shellfish Hatchery supplied us with multiple species of larval and juvenile shellfish for use in experiments.

-1 Figure 7. High frequency (10 min) pHNBS and DO (mg L ) at the Southampton Marine Lab, Easthampton Hatchery, and Islip Hatchery (April – May, 2016).

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Figure 13. Survival (A), growth (B), and development (C) of Argopecten irradians larvae in the diurnal acidification experiment. Percent metamorphosis was calculated 12 days postfertilization. Error bars represent standard deviation of the mean (n = 4). Lowercase letters indicate significant differences.

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Figure 14. Survival (A), growth (B), and development (C) of Mercenaria mercenaria larvae in the diurnal acidification experiment. Percent metamorphosis was calculated 18 days postfertilization. Error bars represent standard deviation of the mean (n = 4). Lowercase letters indicate significant differences.

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Figure 15. Survival (A) and growth (B) of juvenile Argopecten irradians in the diurnal acidification experiment. Error bars represent standard deviation of the mean (n = 4). Lowercase letters indicate significant differences.

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Figure 16. Survival (A) and growth (B) of juvenile Mercenaria mercenaria in the diurnal acidification experiment. Error bars represent standard deviation of the mean (n = 4). Lowercase letters indicate significant differences.

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Figure 17. Survival (A), growth (B), and development (C) of Argopecten irradians larvae in the four treatment diurnal acidification and hypoxia experiment. Percent metamorphosis was calculated 15 days post-fertilization. Error bars represent standard deviation of the

mean (n = 4). Lowercase letters indicate significant differences.

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Figure 18. Survival (A), growth (B), and development (C) of Mercenaria mercenaria larvae in the four treatment diurnal acidification and hypoxia experiment. Percent metamorphosis was calculated 11 days post-fertilization. Error bars represent standard deviation of the mean (n = 4). Lowercase letters indicate significant differences.

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Figure 19. Survival (A), growth (B), and development (C) of Argopecten irradians larvae in the seven treatment diurnal acidification and hypoxia experiment. Percent metamorphosis was calculated 14 days post-fertilization. Error bars represent standard deviation of the mean (n = 4). Both pH and DO negatively affected survival (pH-p < 0.001; DO-p < 0.001), growth (pH-p < 0.001; DO-p < 0.001), and development (pH-p < 0.001; DO-p < 0.001). There was an antagonistic negative effect of both factors on survival (p < 0.05).

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Figure 20. Survival (A), growth (B), and development (C) of Mercenaria mercenaria larvae in the seven treatment diurnal acidification and hypoxia experiment. Percent metamorphosis was calculated 17 days post-fertilization. Error bars represent standard deviation of the mean (n = 4). There were no effects of DO on survival, but pH significantly reduced survival (p < 0.001) and there was an antagonistic negative effect of both pH and DO (p < 0.05). Growth and development of larvae were affected by pH (growth-p < 0.05; development-p < 0.001) and DO (growth-p < 0.05; development-p < 0.001), and there was an antagonistic negative effect of pH and DO on development (p < 0.001).

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Figure 21. Survival (A) and growth (B) of Crassostrea virginica larvae in the seven treatment diurnal acidification and hypoxia experiment. Error bars represent standard deviation of the mean (n = 4). Survival and growth were affected by pH (survival-p < 0.001; growth-p < 0.001) and DO (survival-p < 0.05; growth-p < 0.05) and there was an antagonistic negative effect of pH and DO on growth (p < 0.05).

References Barton, A., Hales, B., Waldbusser, G.G., Langdon, C., Feely, R.A., 2012. The Pacific oyster, Crassostrea gigas, shows negative correlation to naturally elevated carbon dioxide levels: Implications for near-term ocean acidification effects. Limnology and Oceanography 57, 698-710. DePasquale, E., Baumann, H., Gobler, C.J., 2015. Vulnerability of early life stage Northwest Atlantic forage fish to ocean acidification and low oxygen. Marine Ecology Progress Series 523, 145-156. Doney, S.C., Ruckelshaus, M., Duffy, J.E., Barry, J.P., Chan, F., English, C.A., Galindo, H.M., Grebmeier, J.M., Hollowed, A.B., Knowlton, N., Polovina, J., Rabalais, N.N., Sydeman, W.J., Talley, L.D., 2012. Climate Change Impacts on Marine Ecosystems. Annual Review of Marine Science 4, 11-37.

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Gobler, C., DePasquale, E., Griffith, A., Baumann, H., 2014. Hypoxia and Acidification Have Additive and Synergistic Negative Effects on the Growth, Survival, and Metamorphosis of Early Life Stage Bivalves. PLoS One 9.

Gobler, C.J., Talmage, S.C., 2013. Short- and long-term consequences of larval stage exposure to constantly and ephemerally elevated carbon dioxide for marine bivalve populations. Biogeosciences 10, 2241-2253. Green, M.A., Waldbusser, G.G., Reilly, S.L., Emerson, K., O'Donnell, S., 2009. Death by dissolution: sediment saturation state as a mortality factor for juvenile bivalves. Limnology and Oceanography 54, 1037-1047. Talmage, S.C., Gobler, C.J., 2009. The effects of elevated carbon dioxide concentrations on the metamorphosis, size, and survival of larval hard clams (Mercenaria mercenaria), bay scallops (Argopecten irradians), and Eastern oysters (Crassostrea virginica). Limnology and Oceanography 54, 2072-2080. Waldbusser, G.G., Brunner, E.L., Haley, B.A., Hales, B., Langdon, C.J., Prahl, F.G., 2013. A developmental and energetic basis linking larval oyster shell formation to acidification sensitivity. Geophysical Research Letters 40, 2171-2176. Wallace, R.B., Baumann, H., Grear, J.S., Aller, R.C., Gobler, C.J., 2014. Coastal ocean acidification: The other eutrophication problem. Estuarine, Coastal and Shelf Science 148, 1-13.

East Hampton Shellfish Hatchery’s role: provided shellfish and sampling site

Exploring trait-mediated effects of finfish on decapod crustaceans and bay scallops in eelgrass ecosystems

Participants Stephen Heck, Marine Sciences Program, Stony Brook Southampton Bradley Peterson (Advisor)

Summary The bays of Long Island, NY, were once home to vibrant eelgrass meadows, abundant bay scallops (Argopecten irradians), and plentiful fish. Many human activities have altered these ecosystems over the past several decades. Among the plethora of anthropogenic impacts, both commercial and recreational fishing pressure have invariably altered the abundances of large predatory finfish that inhabit the coastal estuaries of Long Island. These include oyster toadfish (Opsanus tau), black sea bass (Centropristis striata), and porgy (Stenotomus chrysops) (Adams 1974, Bigelow and Schroeder 1953). Previous research has demonstrated that these species of finfish prey on several species of crustaceans including black-fingered mud crabs (Dyspanopeus sayi), blue crabs (Callinectes sapidus), and green crabs (Carcinus maenas) (Tettelbach 1986). In turn, these crustaceans are some of the dominant predators of bay scallops. In other ecosystems, large fish predators have been shown to influence the abundance and behavior of crustaceans (Grabowski 2004, McMahan et al. 2013). More specifically these fish both reduce the number of crustacean predators by directly consuming them and lower the feeding rates of the crabs since feeding increases the chance that they will be consumed by fish. We hypothesized that since large fish decrease the abundance of crabs and reduce their feeding rates, the presence of large fish in

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seagrass meadows may indirectly benefit the survival of bay scallops by reducing predation pressure on them.

During August, September, and October of 2016 we set out to explore these hypotheses using large mesocosm tanks at Stony Brook University’s Southampton Marine Station. We also conducted several field experiments during the same months to look at whether the presence of black sea bass and oyster toadfish would indirectly benefit the survival of bay scallops in Shinnecock Bay.

None of our work would have been possible without the help of the East Hampton Town Shellfish Hatchery which provided us with the scallops that we used in our research. Understanding how large finfish may indirectly influence the survival of bay scallops is an important step in figuring out the degree to which different fisheries are connected. Furthermore, this data will be useful in informing cooperative ecosystem-based fisheries management and improving efforts to restore the abundance of the ecologically and economically valuable bay scallop populations.

References Adams, S.M. 1976. Feeding ecology of eelgrass fish communities. Transactions of the American Fisheries Society 105(4):514-519. Bigelow, H.B. and W.C. Schroeder. 1953. Fishes of the Gulf of Maine. Vol. 53. Washington: US Government Printing Office. Grabowski, J.H. 2004. Habitat complexity disrupts predator-prey interactions but the trophic cascade on oyster reefs. Ecology 85(4):995-1004. McMahan, M.D., D.C. Brady, D.F. Cowan, J.H. Grabowski, and G.D. Sherwood. 2013. Using acoustic telemetry to observe the effects of a groundfish predator (Atlantic cod, Gadus morhua) on movement of the American lobster (Homarus americanus). Canadian Journal of Fisheries and Aquatic Science 70(1):1625-1634 Tettelbach, S.T. 1986. Dynamics of crustacean predation on the northern bay scallop, Argopecten irradians irradians. PhD Dissertation. University of Connecticut.

Results: in progress

East Hampton Shellfish Hatchery’s role: provided juvenile bay scallops

Juvenile clam growth and survival in Western Shinnecock Bay Participant Rebecca Kulp, Marine Sciences Program, Stony Brook Southampton

Summary An important part of the Shinnecock Restoration Program is evaluating the ability for juvenile hard clams (Mercenaria mercenaria) to grow and survive in the Weesuck Creek and Tiana Bay clam sanctuaries. Predation mortality in the sanctuary boundaries is an additional concern because the high clam density in clam sanctuary sites (target clam sanctuary density: 25 clams per m2) likely attracts hard clam predators, making it harder for successful clam recruitment. Understanding how juvenile hard clams grow and survive in the current clam sanctuaries will not

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only help to inform whether current clam sanctuaries were placed in an ideal location, but also determine where future clam sanctuary sites should be placed.

Juvenile hard clam growth and survival was compared in and out of three sites: a Weesuck Creek and Tiana Bay clam sanctuary and an eelgrass bed in Tiana Beach. The eelgrass bed site was selected because 1) it is in close proximity to Weesuck Creek and Tiana Bay and 2) eelgrass is a refuge habitat that decreases predation mortality (Wong, 2013). Juvenile hard clams (5-10 mm in shell height [SH]) were individually tagged with bee tags and measured prior to deployment. At Tiana Bay and Tiana Beach, five plots placed in a line 2 m apart were deployed in and out of the clam sanctuary and eelgrass bed. Each plot consisted of 5 tagged hard clams planted inside of a PVC pipe (10 cm in diameter; 6 cm in length) inserted 5 cm into the sediment. By enclosing hard clams with the PVC frame, in-situ clam growth rates could be determined without clams immigrating from the location. To prevent the PVC pipe from sinking in the unconsolidated mud at Tiana Bay, a 2 cm rim of 10 mm Vexar mesh was added at replicates in Tiana Bay. To measure natural clam mortality, an additional protected treatment was added at Weesuck Creek that prevented predator access to hard clams. The protected treatment had a 2 mm window screen mesh covering the top of the PVC pipe. For these treatments, the clams were planted first and then the covered PVC pipe inserted around the clams into the sediment.

Weesuck Creek and Tiana Beach plots were recovered after 5 weeks, whereas Tiana Bay was recovered after 4 weeks. The sediment inside the entire length of the PVC pipe was excavated into a 500 micron mesh bag ensuring clams retained within the PVC pipe were recovered. The number of alive clams recovered per plot was recorded, as well as shell height. Some bee tags fell off of clams, thus a growth rate could not be determined for every clam recovered. Any missing clams were assumed to be consumed in non-protected treatments. Due to differential deployment periods, mortality and growth rates are presented per week.

If the protected treatment plots properly enclosed hard clams without limiting water flow, then there should have always been 100% recovery of clams, alive or dead, with similar growth rates as clams placed in uncovered plots. However, on average there was 2 to 3 clams and a maximum of 5 clams missing from covered plots. Additionally, alive clams recovered from protected treatments grew on average 1 mm less than non-covered treatments and were the only plots recovered with hinged hard clam shells. Perhaps the 2 mm window screen reduced water flow, creating anoxic or suboxic conditions and limiting food delivery. While unlikely there is also the potential that hard clams were able to emigrate out of the bottom of the PVC pipe. Alternatively, with limited visibility at the sites we may have been inconsistent at successfully placing the covered PVC pipes over planted hard clams. Due to the listed potential artifact effects, the rest of results do not include protected plots at Weesuck Creek. Yet, because uncovered PVC pipe plots consistently had clam fragments present-indicative of predation events-we are still considering missing hard clams to have died from predation in uncovered treatments. Therefore, we are also assuming clam emigration from the PVC pipes to be low and equally likely at any of the site plots.

Natural mortality was low to nonexistent at all the sites. Out of the 55 clams recovered only 2 clams died of natural causes; both were initially planted in the eelgrass bed at Tiana Beach. Therefore, predation is the greater concern for newly seeded clams at all sites. The greatest difference in weekly predation mortality (mean ± SD) was at Tiana Beach with 10% ± 8% in the

33 eelgrass bed compared to 21% ± 2.2% outside the eelgrass bed (Figure 1). The clams in the eelgrass bed also had the highest survival out of all the sites, which can be expected due to the high refuge value of eelgrass beds. The high clam densities in the clam sanctuaries did not appear to affect predation, as the difference in mortality was no greater than 4% inside and outside of the clam sanctuaries (Tiana Bay: 16.8% ±. 3.3% in the sanctuary and 12.8% ± 8% outside the sanctuary; Weesuck Creek: 12.8% ± 8.1% in the sanctuary and 13% ± 3% outside the sanctuary).

Within each site, clam growth was similar inside and outside of eelgrass and the clam sanctuaries; weekly growth was less than a 0.1 mm different (Figure 2). This suggests that clam density and the eelgrass structure is not limiting food delivery or altering environmental conditions vital for growth. Weesuck Creek and Tiana Beach had similar growth rates, growing weekly 0.4 ± 0.1 mm and 0.3 ± 0.1 mm, respectively. As indicated by the annual condition index monitoring, Weesuck Creek have been better conditioned than Tiana Bay and may provide ideal conditions for clam growth over Tiana Bay. The maximum clam growth of 4.3 mm over a month occurred at Weesuck Creek. Potentially, Tiana Bay plankton community or bottom environmental conditions does not offer ideal growth conditions. For instance, the bottom sediment in Tiana Bay has high clay content, indicating poor water flow and likely creates stressful conditions for the clams.

Results indicate seeded hard clams should be preferentially planted in the eelgrass beds or Weesuck Creek before Tiana Bay. The eelgrass site had the lowest predation mortality, whereas Tiana Bay had hardly any hard clam growth. By continuing to conduct growth and survival experiments, we can refine our restoration efforts and become closer in reaching our goal of developing sustainable shellfish populations.

Figure 1. The weekly proportion of juvenile hard clam mortality inside and outside of the eelgrass (Tiana Beach) or clam sanctuaries (Tiana Bay and Weesuck Creek).

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Figure 2. The weekly clam growth inside and outside of the eelgrass (Tiana Beach) or clam sanctuaries (Tiana Bay and Weesuck Creek).

References Wong, M.C., 2013. Green crab (Carcinus maenas (Linnaeus, 1758) foraging on soft-shell clams (Mya arenaria Linnaeus, 1758) across seagrass complexity: behavioural mechanisms and a new habitat complexity index. J. Exp. Mar. Biol. Ecol. 446, 139–150. doi:10.1016/j.jembe.2013.05.010

East Hampton Shellfish Hatchery’s role: provided juvenile hard clams

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2016 Public Outreach & Industry Involvement

Outreach & Education

• Barley judged the Montauk Science Fair in January

• East Hampton’s first oyster garden was launched with a pilot program including 15 members, lectures/workshops and a garden in Three Mile Harbor

• Greg Metzger and his Southampton high school marine science class toured the hatchery in February

• Pete discussed shellfish harvesting and the Hatchery’s role in maintaining shellfish populations with East Hampton middle school students in February

• The Dunne Family participated in the Montauk St. Patrick’s Day Parade with a “float” including one of the hatchery trucks and the Sharpie adorned with signage, gear and candy

• Kate participated in the annual Sportsman’s Expo where she presented our work and answered questions from the public

• Barley presented our annual report at the Trustees’ Educational meeting, along with Dr. Gobler’s annual presentation on the state of East Hampton’s waters

• In May we hosted a visitor from Holland’s Stichting Zeeschelp, who was interested in learning more about culturing hard clams

• In June a group of students from the Third House Nature Center toured the hatchery.

• We hosted a Project MOST tour/clamming trip with about 40 kids at the nursery in July

• South Fork Natural History Museum’s summer marine program came to the field site for a tour and oyster seeding in August

• A tour of the field site was conducted for the Ross School summer marine science program in August.

• Montauk Seafood Fest took place in September where Adam represented the hatchery with a touch tank and Kids at our Montauk Seafood Fest displays touch tank

• Barley participated in the annual East Hampton Town Trustees’ Largest Clam Contest presenting our work along with a touch tank and answering questions from the public as well as holding our shellfish counting contest

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• In October, Pete showed a group of Boy Scouts from Sag Harbor around the field site.

• Also in October, two groups of students came to the field site from Hayground School to learn about shellfish farming and participate in seeding.

• The hatchery mentored and provided work space for two 8th grade students from Montauk school as they conducted science experiments utilizing hatchery raised shellfish. These projects will be featured during the Montauk science fair in January. Their abstracts are included in our Projects and Cooperative Research section above.

Committee Involvement

• Barley is chair of the Long Island Shellfish Managers Group

• Barley serves on the New York State Shellfish Advisory Committee

Conference Attendance

• In January, Barley, Kate, Adam, and Carissa attended the Milford Aquaculture Seminar in Connecticut

• Barley attended and displayed a poster about our scallop sanctuary work at the Long Island Natural History Conference in January

Pete giving the Hayground School a tour of the field site

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2016 Infrastructure Management (done in-house unless otherwise noted)

Annual Maintenance • New seals, bearings as necessary, on water pumps • Trailer lights, winches, straps/chains repaired/replaced • All motors winterized • Buoys repainted

Boats/Trailers • New Yamaha outboard on Sharpie • Bilge pump installed in Sharpie • Boats bottom painted • Hulls patched as necessary • Second axle installed on 24’CS trailer

Nursery • Downwelling tanks repaired • Rotten dock decking replaced • Fence repaired and completed on south side

Field Growout • Barge layout reorganized • Ice shield installed on barge for overwintering • Barge pins replaced • New clam bottom bags for over-wintering New setup for sieving post-set oysters • Storage box re-fiberglassed • Five new buoy strings for pearl nets • Old buoy strings overhauled

Montauk Facility • Algae room outlets completed (Parks dept.) • Constructed larger UV/algae transfer hood • New outback tank covers on a pulley system • Constructed set tank ladders • Handles for 10” metal sieves • Mobile touch tank completed New outback tank covers • Decrepit pier/dock dismantled • Re-shingled building entrance • Bathroom walls repaired and repainted, stall removed • Fuji compressor repaired (new fan, bearings, housing) • Saltwater plumbing valves consolidated to one location • New line on intake pulley system • Major de-foliating of overgrown entrance and road (with Parks and Highway Depts.)

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Welding • Aluminum “doohickey” constructed from salvaged material • Backrack for F150 • Davit for 17’Skiff • Cleats under handles on 24’CS • Trailer light protection on flatbed • Mesh bag pressure washing table

Office/Administrative • All years clam seedings tabulated by harbor and site • A lot of headway made on operations manual

Tali driving the skiff loaded with clam blocks

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Appendix I: 2016 Harbor Seeding Maps – All Species

Map 1: Northwest Creek Map 2: Three Mile Harbor Map 3: Hog Creek Map 4: Accabonac Harbor Map 5: Napeague Harbor Map 6: Lake Montauk Map 7: All Harbors

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NORTHWEST CREEK / 2016 SEEDING

CARTOGRAPHY- BARNABY FRIEDMAN 11/15/16 - !( !( !( !(

!( !( !( !(

2013 AERIAL PHOTO

KEY: N

CLAMS: (1 y.o.) 513,704 Avg. Size 11.7mm È NOT TO SCALE Prepared by THE TOWN OF EAST HAMPTON Suffolk County, New York OYSTERS: (1 y.o.) 78,400 Avg. Size 49.9mm

CLAMS OVERWINTERED: (1 y.o.) 850,800 Avg. Size 12.2mm

\\eh-gis-1\DOIT_GIS\Users\Barnaby_Friedman\GIS_Projects\Aquaculture\2016\Northwest Creek_Seeding_2016 THREE MILE HARBOR / 2016 SEEDING CARTOGRAPHY - CARTOGRAPHY BARNABY FRIEDMAN- 11/15/16

2013 AERIAL PHOTO

KEY: N

CLAMS: (1 y.o.) 21,087,965 / (2 y.o.) 157,920 È Avg. Size 6.9mm NOT TO SCALE Prepared by THE TOWN OF EAST HAMPTON Suffolk County, New York OYSTERS: (1 y.o.) 2,031,070 Avg. Size 36.6mm

SCALLOPS: (1 y.o.) 38,250 / (2 y.o.) 31,002 Avg. Size 19mm

\\eh-gis-1\DOIT_GIS\Users\Barnaby_Friedman\GIS_Projects\Aquaculture\2016\ThreeMileHarbor_Seeding_2016 HOG CREEK / 2016 SEEDING CARTOGRAPHY - CARTOGRAPHY BARNABY FRIEDMAN- 11/15/16

2013 AERIAL PHOTO

N KEY: CLAMS: (2 y.o.) 50,400 È Avg. Size 13.8mm NOT TO SCALE Prepared by THE TOWN OF EAST HAMPTON Suffolk County, New York OYSTERS: (1 y.o.) 72,090 Avg. Size 51.8mm

\\eh-gis-1\DOIT_GIS\Users\Barnaby_Friedman\GIS_Projects\Aquaculture\2016\Hog_Creek_Seeding_2016 ACCABONAC HARBOR / 2016 SEEDING CARTOGRAPHY- BARNABY FRIEDMAN 11/15/16 -

2013 AERIAL PHOTO

KEY: N

CLAMS: (1 y.o.) 1,227,840 / (2 y.o.) 121,072 È Avg. Size 11.2mm NOT TO SCALE Prepared by THE TOWN OF EAST HAMPTON Suffolk County, New York OYSTERS: (1 y.o.) 296,724 Avg. Size 49.9mm

CLAMS OVERWINTERED: (1 y.o.) 15,000 Avg. Size 12.5mm

\\eh-gis-1\DOIT_GIS\Users\Barnaby_Friedman\GIS_Projects\Aquaculture\2016\Accabonac_Harbor_Seeding_2016 NAPEAGUE HARBOR / 2016 SEEDING CARTOGRAPHY - CARTOGRAPHY BARNABY FRIEDMAN- 11/15/16

2013 AERIAL PHOTO

KEY: N CLAMS: (1 y.o.) 7,511,038 / (2 y.o.) 121,072 Avg. Size 10mm È CLAMS OVERWINTERED: 30,000 NOT TO SCALE Prepared by Avg. Size 12.5mm THE TOWN OF EAST HAMPTON Suffolk County, New York OYSTERS: (1 y.o.) 901,504 Avg. Size 30.3mm SCALLOPS (2 y.o) 32,909 Avg. Size 35.8mm SCALLOPS OVERWINTERED: 76,800

\\eh-gis-1\DOIT_GIS\Users\Barnaby_Friedman\GIS_Projects\Aquaculture\2016\Napeague_Harbor_Seeding_2016 LAKE MONTAUK / 2016 SEEDING CARTOGRAPHYBARNABY- FRIEDMAN11/15/16 -

2013 AERIAL PHOTO

KEY: N CLAMS: (1 y.o.) 4,566,000 È Avg. Size 6.4mm NOT TO SCALE Prepared by THE TOWN OF EAST HAMPTON Suffolk County, New York OYSTERS: (1 y.o.) 256,818 Avg. Size 52.4mm

\\eh-gis-1\DOIT_GIS\Users\Barnaby_Friedman\GIS_Projects\Aquaculture\2016\Lake_Montauk_Seeding_2016 2016 SEED PLACEMENT N LEGEND CLAMS: 35,357,011 È Not To Scale OYSTERS: 3,636,605 N

SCALLOPS: 102,161 GARDINERS BAY CARTOGRAPHY - CARTOGRAPHY BARNABY FRIEDMAN- 11/16 È Not To Scale

HOG LAKE CREEK MONTAUK

2010 AERIAL PHOTO

THREE MILE NORTHWESTHARBOR HARBOR NAPEAGUE BAY ACCABONAC HARBOR

NORTHWEST CREEK NAPEAGUE HARBOR

2013 AERIAL PHOTO ATLANTIC OCEAN

TOWN OF EAST HAMPTON Prepared By THE TOWN OF EAST HAMPTON Suffolk County, New York Suffolk County, New York

SHELLFISH HATCHERY \\eh-gis-1\DOIT_GIS\Users\Barnaby_Friedman\GIS_Projects\Aquaculture\2016\EH_Town_shellfish_stocking2016

Appendix II: 2016 Harbor Water Temperatures

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PRESENTING East Hampton Town Shellfish Hatchery ANNUAL EVENTS

Open House: In conjunction with Concerned Citizens of Montauk (CCOM) and the South Fork Natural History Museum (SoFo), Hatchery Staff will lead tours through the facility and perform a shellfish spawning demonstration. Typically held in conjunction with the CCOM Earth Day celebration, from 10am to 12pm at the Montauk Shellfish Hatchery.

Oyster Garden Reception: Held at Bay Kitchen Bar in East Hampton in support of the East Hampton Shellfish Education & Enhancement Directive (EHSEED). This is a ticketed event in May: includes beer, wine, oysters, light fare, and a tour of the nursery facility. Please contact SoFo for more information at 631-537-9735.

Town Trustees’ Largest Clam Contest: Submit a clam to be crowned King, or just come for great chowder and clams on the ½ shell. This event usually occurs around the last weekend in September.

Additional events organized by the Hatchery and SoFo are held throughout the year Kids and young adults are welcome to participate! Refreshments will be served & there is no fee for these events! specific dates and information to be announced

Want to schedule your own tour or fieldtrip? Contact us!

WEBSITE: www.ehamptonny.gov/149/aquaculture E-MAIL: [email protected] HATCHERY PHONE: (631) 668-4601 MOBILE PHONE: (631) 816-3082 FOR FURTHER INFORMATION OR TO SIGN UP

These events will go on “rain or shine” – please dress accordingly