Powerhouse Plankton Removers - Marine Biological Laboratory

1
Powerhouse Plankton Removers:
Can shellfish remediate nutrient loading to the coastal pond ecosystems?
Nicholas M. Uline
Gettysburg College
17 December 2013
Advisor: Dr. Ken Foreman, Ecosystems Center, MBL, Woods Hole, MA 02543
2
Abstract
Nutrient over-enrichment from anthropogenic activities can degrade coastal systems,
leading to excessive productions of undesirable primary production and decrease species
diversity in the water column and benthic community. Cultivation of Oysters (Crassostrea
virginica) has been proposed as a remediation strategy that can help improve water quality
through grazing of excess phytoplankton. I assessed the effects of oyster grazing by budgeting
particulate carbon (C) and nitrogen (N) in laboratory microcosms maintained at 20°C. Water
clearance and biodeposition by three size classes of oysters was measured and regressions
relating size to filtration were constructed. Oysters were capable of filtering between 0.5 and 9.5
liters/hr. Oyster respiration and ammonia excretion were all measured. Oysters consumed
between 0.1and 0.5 mg O2/hr and released on average about 0.1 mg ammonia-N per
individual/hr. I used this data together with measured growth rates to evaluate the potential
effect of oyster aquaculture in Little Pond, a 300 hectare embayment in Falmouth MA receiving
approximately 7000 kg of N-loading annually. The model predicts that one million oysters
growing from 5 mm to 64 mm could incorporate about five percent of the load incorporate into
oyster biomass over a 235 day period Direct measurements of the elemental content of oyster
meat and shell in 30 individuals of different sizes reveal that one million oysters growing from 5
to 65 mm in length would incorporate about UU mg N. . These data demonstrate that oysters
have the potential to reverse ecosystem degradation through bivalve-phytoplankton interactions
found in aquaculture.
Introduction
The effects nutrient over-enrichment from land-based human activities has become a
critical issue as society increasingly populates the coastal environment. Eutrophication cause by
anthropogenic activities such as waste water disposal, agriculture practices, urban runoff, and
burning of fossil fuels have degraded more than two-thirds of the coastal rivers and bays in the
United States (Burkholder & Shumway p156). Production of algal biomass, toxic algal blooms,
loss of important near-shore habitat, changes in marine biodiversity and species distribution, and
depletion of dissolved oxygen are potential effects of excess nutrients (Darnell and Rozeat, 1981;
National Research Council 2000). (Carmichael et al. 2012). Understanding the source and
transport mechanisms of particular elements like nitrogen as well as the human activities that
3
alter coastal systems at a local scale are important to developing effective strategies to reduce
negative impacts of eutrophication.
A Massachusetts Estuaries Project (MEP) report(Howes et al. 2006) describes increased
nutrient enrichment particularly in the form of nitrogen (N) loading, to Little Pond, a 300
hectare coastal pond found in the Town of Falmouth, Massachusetts. The primary cause for this
nitrogen-loading is on-site disposal of wastewater and fertilizer from the homes surrounding
Little Pond. (Woods Hole Group 2013). Species that are capable of more rapid nutrient uptake
such as phytoplankton and benthic macroalgae outcompete seagrass (Duarte, 1995; Hein et al.,
1995; Short et al. 1990). Seagrass meadows not only indicate a healthy ecosystem but are an
important role in preserving the environmental quality of estuaries (National Research Council
200) and ultimately provide protection and sediment stability to high species densities and
richness of resident faunas (Orth et al. 1984). Eutrophication has resulted in excess growth of
undesireable macroalgae and loss and degradation of eelgrass beds within Little pond. The
Town of Falmouth is in the process of implementing an approach to reducing the nutrient inputs
into Little Pond with the goal of ultimately restoring eelgrass habitat and desireable benthic
communities .
Oyster cultivation has been proposed as a mitigation measure that also produces valuable
shellfish through sustainable aquaculture since the primary source of nutrition for the oysters is
phytoplankton blooms thatdevelop in response to nutrient loading into coastal ecosystems
(Hargreaves p51). A link between decimation of oyster populations and deteriorating water
quality was proposed by Newell (1988) who suggested that increases in the oyster abundance
could significantly improve water quality. Shellfish, in nature or aquaculture, are able to exert a
“top down” control on phytoplankton concentrations that are supported by enhanced nutrient
inputs (Newell 2004). Previous studies recognize filtration and biodeposition in shellfish
populations as beneficial to water quality and benthic-pelagic coupling, controlling
phytoplankton densities and sequestering nutrients that are removed from the system once the
shellfish are harvested while nitrogen and other nutrients are buried in sediments in the form of
feces and pseudofeces or lost through denitrification (Newell 2004, Kaspar et al. 1985,V erwey
1952). However questions remain about how many oysters are necessary to have the desired
effect on water quality.
4
The most recent alternative approach to manage centralized wastewater processing within
Little Pond is to incorporate juvenile oysters (Crassostrea virginica) within the embayment as an
effort to reduce nutrient-loading and improve overall water quality. In July of 2013,
approximately 1 million juvenile oysters were introduced to Little Pond and cultured in floating
bags, a method that enhances growth rates and reduces mortality to shellfish (Woods Hole Group
2013). Remediation efforts of coastal waters have been taking place in the Chesapeake Bay,
using oyster cultures as a way to clean surface waters that have been enriched from overfertilization, wastewater, and runoff from impervious surfaces (Cerco & Noel 2007). Restoring
bivalve-phytoplankton interactions through shellfish aquaculture has the potential to reverse
ecosystem degradation. Once phytoplankton is removed from the water column, seagrass
meadows are given a greater photosynthetic advantage through the increase of sunlight reaching
the benthos.
In this study I have developed a carbon and nitrogen budget of the oysters used in the
Shellfish Demonstration Project for Little Pond and evaluated their potential to remove nutrients
from the water column as a mitigation measure to improve the health of the embayment. This
includes determining the rate at which oysters graze suspended particulates from the water
column, the amount of nitrogen and carbon they assimilate into their shells and body tissue as
they grow, and finally the total nitrogen oysters export as waste products in the form of feces,
pseudofeces, and ammonium (NH4).
Methods
Grazing Experiments
Grazing rate experiments were run on a total of 12 oysters, 8 from Little Pond and 4 from
Falmouth Harbor. Each oyster was measured for length, width, and thickness using electronic
micrometers. Oysters from three size classes (small oyster from Little Pond, avg. L= 41.99 mm,
W= 29.06 mm, a medium sized oysters from Little Pond,(avg. L= 61.91mm, W= 46.15 mm, and
large oysters from Falmouth Harbor, avg. L= 86.40 mm, W= 52.59 mm were used. Individual
oysters were placed on a platform made from molding clay in a 10 liter aquarium. The clay
platform and oysters were positioned over tin containers that were oriented to separately collect
feces and pseudofeces produced by the oyster. All oysters used the experiments were held in
5
algal free seawater for 24 hours or longer prior to any testing to ensure they cleared their
digestive tracts. Temperature of chambers was maintained at approximately 20 oC at a salinity of
35psu. Each aquarium tank was seton a stir plate with a single stir bar in order to maintain a
uniform phytoplankton mixture. Grazing experiments were run for 10 hours.
Grazing rates were assessed by measuring the clearance of a 50:50 mix of monocultured
phytoplankton includinga flagellate,Tetraselmis chuii, and a diatom ,Chaetoseros calcitrans, to
each aquarium. Aquaria were filled with exactly 8 liters of the phytoplankton mix diluted to a
concentration of between 25-40 µg Chl a L-1 with algal free seawater that was prefiltered through
a 0.45 µm Veraflow Capsule filter. Phytoplankton species were cultured in the Marine
Resource Center (MRC) using f/2 media (Guilliard 1975).. Chlorophyll was measured the stock
cultures to estimate how much of each phytoplankton species should be added to the 0.45 µm
filtered seawater to bring chlorophyll levels within the range of the peak concentrations found in
Little Pond ((25-40 µg Chl L-1,Howes et al. 2006). The amount of stock phytoplankton needed
was then estimated from the proportionality: Cstock Vstock = Ctest concVtest conc
Both spectrophotometric and fluorescence techniques were used to measure chlorophyll
concentrations in the aquaria. Initial and final samples of water (500 ml) from the aquaria were
collected, filtered and extracted in acetone to measure Chlorophyll spectrophotometrically
(Lorenzen 1967). An additional 200 ml water sample was collected at the beginning and end of
each experiment for Particulate Carbon and Nitrogen using a Perkin Elmer 2400 elemental
analyzer. These samples were filtered onto a pre-ashed 25 mm GF/F filter, dried in an oven at
60oC.
In vivo fluorometric measurements of chlorophyll were taken hourly after the
introduction of oysters to the aquaria. The in-vivo estimates of chlorophyll were corrected to
actual Chl a using the spectrophotometer determinations. Based on initial measurements of the
chlorophyll concentration in a 50:50 mix phytoplankton stock solutions, standards were prepared
by diluting to levels of approximately 40, 30, 20, 15, 10, 5, 2 μg L-1. Chlorophyll concentrations
in the standards were then then measured using both in vivo fluorescence and
spectrophotometric methods. The spectrophotometric measurements were assumed to give the
correct and accurate Chlorophyll values and the in vivo measurements were plotted against these
6
values to generate a regression which was used throughout the experiment for converting in vivo
to actual chlorophyll.
Filtration rate K (per individual hour-1) was calculated in vivo fluorescence values taken at the
beginning (x2) and the end (x1) of experimental period during which consistent declines in
chlorophyll occurred (Chlorophyll at time tx > tx+1).
K= (lnx2 – lnx1) / Δt
Volume cleared per hour per individual equals K * volume of the aquarium (8 liters).
Once the final chlorophyll extractions were taken at hour 10, the oysters were allowed to
sit overnight for approximately 12 additional hours to continue expelling any remaining feces
and pseudofeces. The next morning the remaining water was siphoned out and feces and
pseudofeces were collected using a transfer pipette onto a pre-ashed 25 mm GF/F filter.
Biodeposits were dried in an oven at 60oC and stored in small petri dishes. The C and N content
of the biodeposit was determined using a CHN analyzer. The difference in the C and N
concentrations of phytoplankton at the beginning and end of the experiment were used to
determine the total mass of carbon and nitrogen in phytoplankton consumed by each oyster.
Biodeposits were subtracted from this total to understand the nutrients consumed and assimilated
into the tissue and shell.
Dissolved Oxygen (DO) uptake and Ammonium (NH4) release
Approximately 12 hr after each grazing treatment was finished (allowing time for the
oysters to clear all feces and psuedofeces), individuals were transferred into a smaller sealed
chamber filled with 0.45 µm filtered seawater and gently mixed by a magnetic stir bar. The
incubation chambers varied in size based on the size of the oyster. The chamber for the small
oyster contained 1.17 L of algal-free seawater, the medium sized oyster chamber contained 1.59
L, and the largest chamber contained 2.05 L. Oxygen consumption rate was monitored hourly
for 10 hours using a WTW oxygen probe. Twenty .mL of water was collected with replacement
from within each chamber at hours 0, 3, 5, 7, and 10 for NH 4 analysis. Samples were filtered
through GF/F filters, acidified and stored at 4 °C. Ammonia was measured following the
7
protocol of Strickland and Parsons (1972) and based on techniques of the phenol-hypochlorite
method published by Solarzano (1969).
Size Class Assessment
A selection of 25 oysters, 20 from Little Pond and 5 from the Falmouth Harbor, were
selected based on shell length, ranging from 13 mm to 102.50 mm. The Shellfish Demonstration
Project (2013) used Falmouth Harbor oysters as a controlled environmental system to compare
the nursery stock oysters incorporated into Little Pond. All oysters were allowed to filter algalfree seawater for approximately 24 hours before dissected for tissue. This filtering period was to
ensure the oysters cleared all gut content from their digestive tract prior to tissue extraction.
Each shell was hand scrubbed to remove fouling materials and sediment for accurate nutrient
analysis. Both the tissue and dry weight of the shell were measured with an Ohaus Adventurer
balance. Tissues were dried in an oven at 60oC for about 48 hours before being broken up and
packed into a wigglebug to be pulverized into powder for analysis. Shells were allowed to air
dry for 48 hours after being shucked clean of tissues before being pulverized into powder with a
wigglebug. This assessment will provide an estimate of accumulated carbon and nitrogen
assimilated into each size class of oyster. Oyster mass from each size class was then measured
using a CHN analyzer. Shell and dry tissue will provide an estimate of accumulated C and N
biomass as well as an understanding of the projected rate of growth as they mature to harvesting
size (63.5 mm or approx. 2.5 inches and larger in the state of Massachusetts). Based on
measurements and personal communication with Sia Karplus and Chuck Martinsen on the
extraction date from Little Pond, an individual oyster size in the embayment was an average 38
mm (approx. 1.5 in) at the time of extraction and input to Little Pond at an average 5 mm
(approx. 0.20 in).
Results
Individual Oyster Filter-Feeding and chlorophyll a
Phytoplankton abundance determined by extractable chlorophyll (Lorenzen 1967) and
particulate carbon and nitrogen measured in the microcosms at the beginning and end of each
grazing experiment were highly correlated (R2=0.89) and decreased dramatically over the course
of the 10 hour incubation period (Figure 2). The Carbon to Chlorophyll ratio of the
8
phytoplankton was 53.8. Large Oysters (length 86 ± 11 mm) were able graze 92.7% (N=4, SD =
5.7%) of the chlorophyll initially present, while medium sized oysters (61.9 ± 1.4 mm) grazed an
average of 82.8% (N=4, SD=18.7) and the smallest size class (42.0 ± 1.5 mm) grazed 55.9%
(N=4, SD = 10.7%) of the chlorophyll (Figure 3). Although the trends were similar for
particulate C and N, comparison of initial to final particulate concentrations in the water column
suggest that slightly less material was consumed than indicated by the chlorophyll. This may be
due to resuspension of some of the feces and psuedofeces produced during grazing, which could
have contained degraded chlorophyll. Overall, oysters removed 77% of the chlorophyll in the
water column during the incubation period, 66% of the particulate carbon and 56% of the
particulate nitrogen.
In-vivo fluorescence estimates of chlorophyll at the beginning and end of the experiment agreed
well with the extractable chlorophyll and measurements and the standard curve generated from
the dilution of the stock phytoplankton (Extractable Chlorophyll = In vivo chlorophyll/0.1004,
.R2 = 0.97, Figure 4). Using the in-vivo measurements over the period of feeding, I was able to
fit and exponential function to the grazing curves (Figure 5) and obtain a grazing rate estimate
for each oyster (Table 2, Appendix I). Occasionally there was a delay before oysters opened and
fed, but the delay was independent of the concentration of phytoplankton at the initial time of the
treatment. Clearance rates were calculated for each oyster only for the period during which the
oysters actually filter-fed. The largest oysters were able to deplete phytoplankton in the
experimental chamber to a minimum of 2.26 μg Chl a L-1. Smaller oysters, however, over the
same amount of time could only deplete chlorophyll a levels to about 9.34 μg L-1 (Table 2). .
Plotting the clearance rate estimates for each oyster against the length of the shell, I was able to
fit a power function (Y=0.0003X2.1808, N= 12, R2 = 0.63) that predicts the size dependent
filtration rate for any given length of an individual (Figure 6). Filtration rates increased about
seven fold over the range of sizes found for the oysters (a 40 mm long oyster would filter about
0.93 liters/hr while a 100 mm long oyster would clear about 6.9 liters/hr).
Respiration, Ammonia Excretion and Carbon and Nitrogen Budgets
Roughly 12 hours after each feeding experiment was completed and the oysters had cleared their
digestive tract of feces and psuedofeces, an estimate of net growth was obtained by measuring
excretion and respiration by transferring each oyster to a sealed chamber and measuring
9
ammonia release and oxygen uptake over a 10 hr period. The oysters showed linear rates of
oxygen consumption and ammonia production (Figures 7 and 8) and were correlated with size of
the individual (Figure 9). Oxygen uptake during respiration was converted to CO2 release
assuming an RQ of one.
Carbon and nitrogen budgets based on this data are compiled for individual oysters in Table 1
and shown graphically in figure 10. Consumption was estimated as initial – final particulate C
and N in the grazing experiments. Egestion was measured as feces and pseudofeces collected 12
hours after grazing experiment ended to allow clearing of the oysters gut. Losses from
respiration and excretion were also measured. Estimates of net assimilation (consumption –
egestion) and growth (assimilation – respiration or excretion) were scaled for a 10 hour period,
which was the duration of the grazing experiments, and are shown for each oyster across the size
spectrum (97 - 40 mm in length) in Table 1. Oysters that did not produce biodeposits over the
~12 post experiment period were excluded in determining assimilation efficiency by each oyster.
On average, oysters ingested about 11.67 mg of C and 1.51 mg of N during the grazing
experiments. About 34.5% of the phytoplankton carbon and 47.3% of the nitrogen consumed
was egested as either feces or psuedofeces (Figure 11). In contrast to filtration rate, there was no
relationship between size and egestion; on average oysters generated about 0.4± 0.06 mg C and
1
0.07 ± 0.001 mg N per hour of biodeposits. Assimilation efficiencies averaged 63.6 ± 6.9%
(mean and SE) for C and 47.4 ±12.1% for N (Table 1). Net growth efficiency was about 85%.
For nitrogen, assimilation efficiency was about 52.2% with a net growth efficiency of 87.5%.
Summarizing the average fate of carbon consumed by oysters across all size classes measured, it
appears that about 7 % of C and -9% of N in phytoplankton ingested was released in feces, 27 %
of C and 38% of N was released in pseudofeces and 10% of the assimilated C was respired while
about 7% of the assimilated N was excreted (Figure 11),
Size Class Assessment - Nitrogen and Carbon Accumulation during Growth
Both shell and meat mass increased exponentially when plotted against length (R 2=0.97 and
0.94, respectively). The dry weight of the shell mass on average was about 13 times greater than
dry weight of the meat. The molar C:N ratio of the meat increased as the animals grew from 4.7
1
Mean ± standard error (N = 12)
10
in the smallest individuals to 7.2 in the largest, suggesting greater storage of carbon rich,
nitrogen poor energy compounds like glycogen and lipids in their tissues (Figure 12, top graph).
On average for the 25 individuals measured, the molar C:N ratio in the meat was about 5.36
while the average C:N in shell was 89. Nitrogen content could be accurately estimated from the
length using an exponential function (mg N/individual = 0.0004*Length (mm)2.9164, R2=0.96,
Figure 12, bottom graph). Oysters accumulated a total of about 290 mg N/individual as they
grew from 13 mm to 102 mm in length, and about half this N was in meat and half in shell.
Discussion
My goal is to use data collected in the lab to estimate filtration and nutrient removal by
oyster populations being cultured in the field, specifically in Little Pond. A number of studies
have used calculations and extrapolations from laboratory analyses to estimate grazing and
biodeposition by filter feeders in the field (Doering & Oviatt 1986, Grizzle et al. 2006). I
constructed a model using an average chlorophyll a values observed in Little Pond and total N
loading estimated for Little Pond (Howes 2006, Table 5). The model incorporated the filtration
rate and the average assimilation efficiency of N results from the grazing experiments reported
here. A linear rate of growth (mm day-1) was assumed for the growth of the oysters in the
Shellfish Demonstration Project, which increased in size from the day they were placed into the
system at 5 mm length (July 4th, 2013) to 38 mm length on the day they were removed from the
system (November 13th, 2013). The total consumption of N was then summed over a period of
235 days that was estimated for the oysters to grow to market size (63.5 mm). The model
included the sum of N removal from Little Pond, addressing the 1 million oysters originally
incorporated over this time period (Figure 13). The model assumes that N that would be only be
assimilated to the body tissues and disregards any N that would be transferred back into the
water column as excreted waste or to the sediment surface as biodeposits. Based on the filter
feeding, assimilation and net growth rates, the model predicts 5.09% of total annual N inputs to
Little Pond will be taken up by oysters. The model also includes the total N removal of 5 million
oysters to show the necessary capacity that must be met in order to remove over 25% of the N
inputs to Little Pond.
An additional alternative model was constructed using the power equation that was
calculated after determining the N content within the total dry weight in biomass of whole
11
oysters in relation to length (Figure 12). This alternative approach estimates that only 1.13%
(43.45 kg N) of the total N input to Little Pond would be removed with 1 million oysters once
market size length was reached. A total of 5.64% (392.41 kg N) from the same N load would be
removed if the Shellfish Demonstration Project included 5 million oysters.
These two independent approaches to estimating N-sequestration in oyster populations
show that cultivation and harvesting of one million oysters in Little Pond could remove between
1 and 5% of the total loading to the pond.
My data and the model suggests that oysters have the potential to reverse ecosystem
degradation through bivalve-phytoplankton interactions found in aquaculture. Oysters of all size
classes used in the laboratory experiment were capable of depleting phytoplankton concentrations at an exponential rate. My study shows that oysters indeed exploit as a resource the
intensive phytoplankton concentrations within the chamber, which in field studies may otherwise
be viewed by water quality managers as a nuisance (Lindahl et al. 2005). Implementing bivalve
aquaculture in eutrophic coastal waters has been considered as a means of restoring an impacted
ecosystem, clearing suspended particulates from the water column (Nelson et al. 2004).
The negative impact of undesired phytoplankton blooms includes the shading of
submerged aquatic vegetation. As decaying phytoplankton sink to the sediment, they stimulate
bacterial respiration that can ultimately lead to benthic hypoxia and anoxia. A previous report on
the water quality of Little Pond occasionally found benthic communities to have oxygen level
below 3 mg L-1 (Howes et al. 2006). This level creates stressful conditions for the inhabiting fish
species as well as the benthic communities. The details of benthic-pelagic coupling between
bivalves filter-feeding phytoplankton and their environment are not completely understood, but
data from other studies supports my laboratory results of the significant impact aquaculture
farms have on phytoplankton communities on a system-wide scale (Dame 1996, Pietros and Rice
2003). Seagrass bed restoration efforts are a key component to the Shellfish Demonstration
Project and by removing phytoplankton and other particulate organic matter from the water
column, a photosynthetic advantage to the seagrass beds is provided with increases in light
penetration. Field studies on aquaculture in the Chesapeake Bay and models on dense oyster
populations appear to enhance seagrass abundances and production via a positive feedback loop
(Grizzle et al. 2006, Cerco & Noel 2007).
12
My study shows that about half the phytoplankton biomass that is grazed by oysters is
transferred as biodeposits to the sediment surface. The assimilation efficiencies of C and N by
individual oysters varies widely, however averages of assimilated material are supported by
other estimates of bivalves to digest and absorb about 50% of the particulate N that they filter
from the water column (Newell and Jordan 1983). While this study addressed the differences in
clearance rate as beneficiary of filter-feeders, an additional consideration must account for the
amount of nutrient reintroduced to the system as waste products in the form of feces,
pseudofeces, ammonium, and carbon dioxide. A study by Bayne and Newell (1983) described a
so-called “pseudofeces threshold for bivalves, showing a portion of the particles captured are
rejected before ingestion because respiratory activities of the gills take precedence over food
capture (Wikfors p128). This release of excessive biodeposits would be expected to have a
direct effect on the ecosystem, thus the concept of “carrying capacity” for oyster aquaculture
must be considered spatially and defined by the limit of resources available.
The filter-feeding activities of cultured bivalves can be viewed as a natural solution to the
current problem in Little Pond. Sedimentation of biodeposits has been found to have few effects
on native macrobenthos (Dumbauld et al. 2009, Fabi et al. 2009) as well as capabilities of
mussels and oysters to potentially enhance epifaunal diversity (Commito et al. 2008). For future
projections of the Shellfish Demonstration Project, comparisons of the benthic community,
including changes to the water column above the sediment, must be further assessed before and
after the incorporation of oysters. Future in situ methods should also consider environmental
factors such as temperature, salinity, growth rate and seasonality during the course of
investigation given that these variables remain constant in this laboratory experiment.
Acknowledgements
Funding and support was provided by the Semester in Environmental Science program at
the Marine Biological Laboratories in Woods Hole, Massachusetts. I thank Ken Foreman, Scott
Lindell, Rich McHorney, Alice Carter, Fiona Jevon, Sarah Nalven, Patrick Doughty, Sia
Karplus, and Chuck Martinsen for assistance with logistics, field site selection, coordination, and
laboratory analysis.
13
Literature Cited
Bayne, B.L. and R.C. Newell. 1983. Physiological energetics of marine molluscs. The Mollusca
4: 407-515.
Burkholder, JoAnn M. and Sandra E. Shumway. 2011. Bivalve shellfish aquaculture and
eutrophication. Pages 155-215 in Shumway, Sandra E., editor. Shellfish Aquaculture and
the Environment. West Sussex: John Wiley & Sons, Inc.
Carmichael, Ruth H., A.C. Shriver and I. Valiela. 2012. Bivalve response to estuarine
eutrophication: the balance between enhanced food supply and habitat alterations.
Journal of Shellfish Research 31:1-11.
Cerco, Carl F. and Mark R. Noel. 2007. Can oyster restoration reverse cultural eutrophication in
Chesapeake Bay? Estuaries and Coasts 30:331-343.
Commito, J.A., et al. 2008. Species diversity in the soft-bottom intertidal zone: biogenic
structure, sediment, and macrofauna across mussel bed spatial scales. Journal of
Experimental Marine Biology and Ecology 366: 70-81.
Council, National Research. 2000. Clean Coastal Waters: Understanding and Reducing the
Effects of Nutrient Pollution. National Academy Press ,Washington D.C.
Dame, R.F. and T.C. Prins. 1998. Bivalve carrying capacity in coastal ecosystems. Aquatic
Ecology 31: 409-421.
Darnell, Rezneat M., and T.M. Soniat. 1981. Nutrient enrichment and estuarine health. Pages
225-245 in B.J. Neilson, and L.E. Cronin, editors. Estuaries and Nutrients. Humana
Press, Clifton, New Jersey, USA.
Doering, Peter H. and Candace A. Oviatt. 1986. Application of filtration rate models to field
populations of bivalves: an assessment using experimental mesocosms. Marine Ecology
31: 265-275.
Duarte, C.M. 1995. Submerged aquatic vegetation in relation to different nutrient regimes.
Ophelia 41: 87- 112.
Dumbauld, B.R., J.L. Ruesink and S.S. Rumrill. 2009. The ecological role of bivalve shellfish
aquaculture in the estuarine environment: a review with application to oyster and clam
culture in West Coast (USA) estuaries. Aquaculture 290: 196-223.
Fabi, G., S. Manoukian and A. Spagnolo. 2009. Impact of an open- sea suspended mussel
culture on macrobenthic community (Western Adriatic Sea). Aquaculture 289: 54-63.
14
Group, Woods Hole. 2013. Final Shellfish Stock Assessment Report of Findings - Little Pond,
Falmouth, Massachusetts. Falmouth.
Grizzle, Raymond E., et al. 2006. A new in situ method for measuring seston uptake by
suspension-feeding bivalve molluscs. Journal of Shellfish Research 25: 643-649.
Hargreaves, John A. 2011. Molluscan shellfish aquaculture and best management practices.
Pages 51-80 in Shumway, Sandra E., editor. Shellfish Aquaculture and the Environment.
West Sussex: John Wiley & Sons Inc.
Hein, M., M.F. Pedersen, and K. Sand-Jensen. 1995. Size-dependent nitrogen uptake in microand macroalgae. Marine Ecology Progress Series. 118: 247-253.
Howes, B.L., et al. Linked Watershed-Embayment Model to Determine Critical Nitrogen
Loading Thresholds for the Little Pond System, Falmouth, MA. Boston: SMAST/DEP
Massachusetts Esuaries Project, Massachusetts Department of Environmental Protection,
2006.
Karplus, Sia. Personal communication.
Kaspar, H.F., et al. 1985. Effects of mussel aquaculture on the nitrogen cycle and benthic
communities of Kenepuru Sound, Marlborough Sounds, New Zealand. Marine Biology
85:127-136.
Lorenzen, C.J. 1967. Determination of chlorophyll and pheo-pigments: spectrophotometric
equations. Limnology and Oceanography 12:343-346.
Lindahl, O., R. Hart and B. Hernoth. 2005. Improving marine water quality by mussel farming:
a profitable solution for Swedish society. Ambio 34: 131-183.
Martinsen, Chuck. Personal communication.
Nelson, K.A., et al. 2004. Using transplanted oyster (Crassostrea virginica) beds to improve
water quality in small tidal creeks: a pilot study. Journal of Experimental Marine
Biology and Ecology 298: 347-368.
Newell, R.I. and S.J. Jordan. 1983. Preferential ingestion of organic material by the American
oyster, Crassostrea virginica. Marine Ecology Progress Series 13: 47-53.
Newell, Roger. 1988. Ecological changes in the Chesapeake Bay: are they the result of
overharvesting the American oyster, Crassostrea virginica. Understanding the Estuary:
Advances in Chesapeake Bay Research 29-31.
15
Orth, Robert J., Kenneth L. Heck and Jacques van Montfrans. 1984. Faunal communities in
seagrass beds: a review ofthe influence of plant structure and prey characteristics on
predator: prey relationships. Estuaries 339-350.
Pietros, J.M. and M.A. Rice. 2003. The impacts of aquacultured oysters, Crassostrea virginica
(Gmelin, 1791) on water column nitrogen and sedimentation: results of a mesocosm
study. Aquaculture 220: 407-422.
Short, F.T., W.C. Dennison, and D.G. Capone. 1990. Phosphorous-limited growth of the
tropical seagrass Syringodium filiforme in carbonate sediments. Marine Ecology
Progress Series 62: 169-174.
Solarzano, L. 1969. Determination of ammonium in natural waters by phenol hypochlorite
method. Limnology Oceanography 14:799-800.
Strickland, J.D.H. and T.R. Parsons. 1972. A practical handbook of Seawater Analysis.
Fisheries Research Board of Canada.
Verwey, J. 1952. On the ecology and distribution of cockle and mussel in the Dutch Wadden
Sea, their role in sedimentation and the source of their food supply. Archives
Neerlandaises Zoologie 10:171-339.
Wikfors, Gary H. 2011. Trophic interactions between phytoplankton and bivalve aquaculture.
Pages 125-134 in Shumway, Sandra E., editor. Shellfish Aquaculture and the
Environment. West Sussex: John Wiley & Sons, Inc.
16
Tables And Figures
Figure 1: The location of the study site used the Shellfish Demonstration Project between July 4thto
November 13th 2013. The project incorporated roughly 1 million oysters in floating cages in this
particular area.
17
3000
y = 53.844x + 419
R² = 0.8968
Carbon (µg/liter)
2500
2000
1500
1000
End of Experiment
Beginning of Experiment
500
0
0
10
20
30
40
50
Chlorophyll (µg/liter)
500
y = 7.4802x + 94.368
R² = 0.8945
Nitrogen (µg/liter)
400
300
200
100
0
0
10
20
30
40
50
Chlorophyll (µg/liter)
Figure 2: Relationship between carbon and chlorophyll (top panel) and nitrogen and chlorophyll (bottom
panel) in particulate matter in the aquaria used in grazing experiment at the beginning of the
experiments (dark blue symbols) and end (green symbols). Different phytoplankton concentrations
were used during each day the experiments were conducted and initial concentrations ranged from 25
to 40 µg Chl/L.
18
Figure 3. Average percentage of total particulate material added to 10 liter aquaria that was grazed by
large, medium and small oysters during feeding trials expressed as chlorophyll, carbon and nitrogen.
Fluorometer (μg L-1)
19
7
6
5
4
3
2
1
0
y = 0.1004x
R² = 0.9753
0
20
40
Spectrophotometer (μg L-1)
60
80
Figure 4: The dilution (estimated at 60, 45, 30, 15, 5, and 2 micrograms chlorophyll a per liter) standard
curve of chlorophyll a (green dots) measured via spectrophotometer using the Lorenzen (1976) method
in relationship to in vivo fluorescence using a fluorometer, resulting in the linear equation shown in
green. Chlorophyll a concentrations before and after each individual filtering experiment (red asterisks)
were applied to the graph to show accuracy of fluormetric readings taken during the experiment.
Little Pond (40.09mm)
[Chlorophyll a] (ug L-1)
40.0
Little Pond (62.85mm)
Falmouth Harbor (69.15mm)
35.0
30.0
25.0
20.0
15.0
10.0
5.0
0.0
0
2
4
Time (Hours)
6
8
Figure 5: An example of in vivo fluorescence in experiments with cultured phytoplankton
concentrations (micrograms of chlorophyll a per liter) over filtration time (hours). Notice the difference
in scale across size class.
10
20
Filtration Rate (L indiv. -1 hr-1)
10
8
y = 0.0003x2.1808
R² = 0.6343
6
4
2
0
40
50
60
70
Length (mm)
80
90
100
Figure 6: The size dependent filtration rate (liters per individual per hour) in relationship to length
(millimeters) of experimental oysters represented as a power function. This equation is applied to the
model of Little Pond.
21
Oxygen (mg L-1)
11
9
Falmouth (87.09mm)
Little Pond (61.37mm)
y = -0.3251x + 7.9878
R² = 0.6363
y = -0.4875x + 8.4288
R² = 0.8244
Falmouth (98.60mm)
Little Pond (59.86mm)
y = -0.2488x + 6.2568
R² = 0.924
y = -0.175x + 6.1583
R² = 0.8962
Little Pond (43.61mm)
y = -0.2227x + 7.728
R² = 0.8732
7
5
Oxygen (mg L-1)
3
9
7
y = -0.114x + 6.1861
R² = 0.8883
5
3
9
Oxygen (mg L-1)
Little Pond (40.88mm)
Falmouth (92.57mm)
y = -0.1313x + 5.7696
R² = 0.7364
7
Little Pond (63.54mm)
Little Pond (43.37mm)
y = -0.1623x + 5.9708
R² = 0.8802
y = -0.0996x + 5.8786
R² = 0.82
5
Oxygen (mg L-1)
3
9
7
Falmouth (69.15mm)
Little Pond (62.85mm)
Little Pond (40.09mm)
y = -0.1422x + 5.7699
R² = 0.9337
y = -0.1302x + 5.7897
R² = 0.9396
y = -0.0524x + 5.7772
R² = 0.6959
5
3
0
2
4
6
Time (Hours)
8
10
Figure 7: Oxygen consumption (milligrams per liter) during incubation experiments over time
(hours) with various size classes of oysters.
22
mg N/individual
0.60
Falmouth (87.09mm)
Little Pond (61.37mm)
y = 0.0066x + 0.0884
R² = 0.4621
0.40
y = 0.0308x + 0.1225
R² = 0.9393
Little Pond (43.61mm)
y = 0.0099x + 0.019
R² = 0.9369
0.20
0.00
Falmouth (98.60mm)
mg N/individual
0.20
mg N/individual
Little Pond (40.88mm)
y = 0.0144x + 0.0546
R² = 0.7803
0.10
y = 0.0106x + 0.0416
R² = 0.8718
y = 0.0053x + 0.0553
R² = 0.7467
0.00
Falmouth (92.57mm)
mg N/individual
Little Pond (59.86mm)
0.20
Little Pond (63.54mm)
Little Pond (43.37mm)
y = 0.0115x + 0.0514
R² = 0.9713
0.10
y = 0.0072x + 0.105
R² = 0.7253
y = 0.0115x + 0.0449
R² = 0.992
0.00
0.15
Little Pond (40.09mm)
0.10
y = 0.0021x + 0.0313
R² = 0.6578
Little Pond (62.85mm)
y = 0.0079x + 0.0107
R² = 0.906
Falmouth (69.15mm)
y = 0.007x + 0.0149
R² = 0.9024
0.05
0.00
0
2
4
Time (Hour)
6
8
Figure 8: Ammonium excretion (milligrams NH4 per individual) in incubation experiments over time
(hours) using three different size classes of oysters.
10
mg O2/ individual/hour
23
0.70
0.60
0.50
0.40
0.30
0.20
0.10
0.00
y = 0.0064x - 0.1316
R² = 0.6121
40
50
60
70
Length (mm)
80
90
100
Figure 9: The relationship between oxygen consumption (milligram O2 per individual per hour) and
length (millimeter) of each oyster used in the chamber experiment observing respiration and excretion
of oysters over an allotted period of time. This shows a linear rate of oxygen consumption given the size
of the particular oyster.
24
Figure 10: Fates of carbon and nitrogen in oysters of different sizes measured a the mass found in feces,
psuedofeces, or consumed in respiration during 10 hour incubations. The difference between the total
mass consumed and the sum of feces, pseudofeces and respiraton was assumed to equal the amount of
N or C incorporated for growth.
25
Figure 11: Average fate of C and N in phytoplankton consumed by oysters showing amounts allocated
toward growth, or released as feces, pseudofeces, respiration and excretion. Labels for each pie section
give mean mass (mg /10 hr) and percent of total consumption averaged for all oysters across the size
spectrum (40 mm to 97 mm).
26
8.0
Molar C:N Ratio
7.0
6.0
5.0
y = -0.4098x2 + 2.1008x + 4.5555
R² = 0.932
4.0
3.0
0.0
0.5
1.0
1.5
2.0
2.5
3.0
3.5
Tissue Dry Weight (mg)
Dry Weight (mg N)
350
Whole Oyster
300
y = 0.0004x2.9165
R² = 0.9617
250
Tissue
Shell
y = 0.0005x2.8006 y = 0.0179x2 - 0.6948x
R² = 0.9488
R² = 0.9388
200
150
100
50
0
0
20
40
60
80
100
Length (mm)
Figure 12: The relationship between the dry weight (milligrams of nitrogen) and length (millimeters) of
the 25 oysters selected for nutrient analysis. Tissue material corresponds to a polynomial function while
shell material from each oyster is represented by a corresponding power function. Their total sum
represents the total dry weight of N found in the whole oyster which is also shown by a power function
according to length. See Table 3 in Appendix I for actual values.
27
N Assimilation (kg)
25
20
15
10
1 Million Oysters
5 Million Oysters
5
0
0
50
100
150
200
Days in Little Pond
Figure 13: Model estimate of N assimilation by a population of one and five million oysters growing
from five mm to market size (63.5 mm) over 235 days . N-assimilation represents the total amount of N
that would be removed from Little Pond if the oysters were able to grow to market size (63.5 millimeters) over a 235 day period given an assumed linear growth rate from the Shellfish Demonstration
Project (5 – 38 millimeters on average) and an average chlorophyll a concentration from the MEP Report
(2006).
28
Table 1: The total mass of of carbon (C) and nitrogen (N) consumed (milligram/10 hrs) for all individual oysters ranging from 96.8 to 40.09 mm in
length. Also shown is the mass of biodeposits (feces and pseudofeces) produced, the amount of C assimilated that was lost through respiration
of CO2 estimated from oxygen uptake and the amount of N lost through excretion from ammonium release. Assimilation and Net Growth over
the 10 hr incubation period was estimated and both assimilation efficiency and net growth efficiency was calculated. Average values are given in
the right hand column.
Length (mm)
96.8
92.6
87.1
69.2
63.5
62.9
61.4
59.9
43.6
43.4
40.9
40.1
Mean
Consumption of C (mg)
12.04
12.21
12.87
17.49
13.3
15.93
8.07
10.51
11.73
7.24
9.29
9.36
11.67
Feces C (mg/10 hr)
Pseudo C (mg /10hr)
0.36
3.60
0.72
3.98
0.62
1.01
0.20
2.95
1.16
5.30
1.44
4.91
0
0
1.01
4.80
0.26
0.71
1.76
4.20
0.73
5.02
1.83
1.80
0.84
3.19
Total C Egestion (mg/10 hr)
3.96
4.70
1.63
3.15
6.46
6.35
0
5.81
0.97
5.96
5.75
3.63
4.03
8.08
7.52
11.24
14.35
6.84
9.58
8.07
4.70
10.76
1.28
3.54
5.73
7.64
1.91
1.01
2.50
1.09
0.97
0.78
2.91
1.04
0.98
0.44
0.50
0.23
1.20
6.17
6.51
8.74
13.25
5.88
8.80
5.16
3.66
9.78
0.85
3.04
5.50
6.44
67.1%
61.5%
87.3%
82.0%
51.5%
60.1%
100.0%
44.7%
91.7%
17.7%
38.1%
61.2%
65.5%
76.3%
86.6%
77.8%
92.4%
85.9%
91.9%
64.0%
77.8%
90.9%
65.9%
85.9%
96.0%
84.3%
Consumption of N (mg)
1.71
1.67
1.44
2.55
1.98
2.23
0.79
1.43
1.37
0.73
1.18
1.06
1.51
Feces N (mg)
Pseudo N (mg)
0.056
0.629
0.112
0.697
0.109
0.169
0.032
0.479
0.189
0.950
0.241
0.886
0
0
0.163
0.866
0.043
0.114
0.293
0.834
0.113
0.970
0.296
0.331
0.137
0.577
Total N Egestion
0.685
0.809
0.278
0.511
1.139
1.127
0
1.029
0.157
1.127
1.083
0.627
0.714
1.025
0.861
1.162
2.039
0.841
1.103
0.79
0.308
1.213
-0.397
0.097
0.433
0.790
0.144
0.115
0.000
0.070
0.072
0.079
0.308
0.106
0.099
0.115
0.053
0.021
0.099
Net Growth
(Assimilation - Excretion)
0.881
0.746
1.162
1.969
0.769
1.024
0.482
0.202
1.114
-0.512
0.044
0.412
0.691
Assimilation efficiency for N
59.9%
51.6%
80.7%
80.0%
42.5%
49.5%
100.0%
21.5%
88.5%
-54.4%
8.2%
40.8%
52.2%
Net Growth Efficiency for N
86.0%
86.6%
100.0%
96.6%
91.4%
92.8%
61.0%
65.6%
91.8%
129.0%
45.4%
95.2%
87.5%
Assimilation
(Consumption - Egestion)
Respiration (mg C / 10 hr)*
Net Growth (mg/10 hr)
(=Assimilation - Respiration)
Assimilation Eff. for C
(Assim/consumption)
Net Growth Efficiency for C
(=Growth/Assimilation)
Assimilation
(Consumption - Egestion)
Excretion (mg N)
*From oxygen assuming RQ of one.
29
APPENDIX I (Tabular Data)
Table 2: Individual oysters from Falmouth Harbor (FH oyster) and Little Pond (LP large or LP small) used
in filtering experiment represented in order according to length (millimeters) and the in vivo
fluorescence concentrations (microgram chlorophyll a per liter) used in the equation modified from A.E.
Thessen et al. (2010) over the change in time (Δt), providing a rate (K) for filtration.
Location
Length
(mm)
Falmouth Harbor
Falmouth Harbor
Falmouth Harbor
Falmouth Harbor
Little Pond
Little Pond
Little Pond
Little Pond
Little Pond
Little Pond
Little Pond
Little Pond
98.60
92.57
87.09
69.15
63.54
62.85
61.37
59.86
43.61
43.37
40.88
40.09
Tissue
(g dry
wt.)
2.501
2.071
1.725
0.866
0.672
0.650
0.606
0.562
0.218
0.214
0.180
0.170
Initial
ug
chla/L
24.3
40.93
28.36
36.54
40.93
36.54
28.36
24.3
28.36
40.93
24.30
36.54
Δt
Final ug
chla/L
(hour)
0.375
0.1
2.26
1.31
0.25
4.86
12.68
1.31
9.71
9.65
9.34
21.49
8.13
5.03
6.03
9.05
9.15
10.00
10.00
9.20
8.07
7.10
9.20
7.93
Rate (K)
Volume
L/indiv/hour
0.513
1.196
0.419
0.368
0.557
0.202
0.0804
0.317
0.133
0.204
0.104
0.067
4.105
9.566
3.356
2.942
4.457
1.614
0.644
2.540
1.063
1.628
0.831
0.536
30
Table 3: The total number of oysters used for shell and tissue extraction to determine nutrient content
in relation to size (20 from Little Pond and 5, in red, from Falmouth Harbor). These numbers were used
to construct the power function in Figures 12.
Oyster
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
Length
(mm)
13.24
16.32
17.05
18.71
21.81
23.35
25.3
27.42
31.55
32.28
33.89
37.11
39.21
43.16
44.04
50.13
56.02
58.45
62.67
72.61
73.03
78.11
82.24
95.35
102.5
Size (mm)
Width
(mm)
11.85
11.16
13.23
15.48
15.84
16.25
18.00
16.18
19.78
19.02
22.79
25.44
25.15
31.64
33.29
36.06
44.46
50.35
44.96
47.60
51.42
59.12
44.57
58.52
56.50
Dry Weight (g)
Thickness
(mm)
4.33
4.13
4.84
4.75
5.05
4.87
4.70
7.61
5.65
8.15
8.39
10.18
9.66
12.34
15.16
9.71
14.64
15.43
17.21
17.34
17.13
26.39
26.14
24.18
31.44
Tissue
0.019
0.015
0.016
0.019
0.022
0.030
0.031
0.036
0.031
0.055
0.061
0.096
0.132
0.252
0.328
0.457
0.410
0.640
0.667
0.609
1.799
2.944
1.554
2.069
2.803
Shell
0.163
0.133
0.184
0.238
0.285
0.464
0.636
0.664
0.876
0.997
1.216
1.664
2.056
3.785
3.804
5.150
6.793
7.115
8.087
8.909
20.901
42.793
32.655
37.825
66.563
Tissue
%C
36.46%
39.36%
42.25%
39.70%
37.14%
37.72%
38.30%
38.88%
39.46%
40.79%
42.12%
40.75%
39.38%
39.51%
39.63%
40.12%
40.60%
40.95%
41.30%
42.85%
44.40%
42.93%
41.45%
41.85%
42.25%
%N
9.13%
9.84%
10.55%
10.03%
9.50%
9.59%
9.67%
9.76%
9.84%
10.14%
10.43%
10.03%
9.62%
9.15%
8.67%
9.10%
9.53%
9.27%
9.01%
8.14%
7.26%
7.07%
6.88%
6.85%
6.81%
Shell
%C
11.96%
11.99%
12.02%
12.07%
12.11%
12.08%
12.05%
12.03%
12.00%
12.04%
12.07%
12.05%
12.02%
12.05%
12.08%
11.71%
11.33%
11.76%
12.18%
12.17%
12.15%
12.14%
12.12%
12.38%
12.64%
%N
0.13%
0.14%
0.15%
0.17%
0.19%
0.17%
0.14%
0.15%
0.15%
0.16%
0.16%
0.15%
0.13%
0.14%
0.15%
0.15%
0.14%
0.16%
0.18%
0.19%
0.19%
0.18%
0.17%
0.19%
0.21%
31
Table4: The modelled figure of the total nitrogen removed from Little Pond, comparing 1 million oysters
and 5 million, was designed using an average rate of growth given from the Shellfish Demonstration
Model (Input July 4, 2013 and output November 13, 2013). The model uses the filtration rate and
assimilation efficiency results from the experiment and applied over a 235 day period (the total number
of days estimated to reach market size).
Shellfish Demonstration Project Model
Average Growth (mm d-1)
0.254
Total Consumption
(kg N 1million indiv-1)
794.8
Biodeposition Rate
(kg N 1million indiv-1)
445.1
(
Assimilated N
(kg N 1million indiv-1)
349.7 (