THE EFFECT OF SALINITY ON EASTERN OYSTER REPRODUCTION IN THE HUDSON RIVER ESTUARY
A Final Report of the Tibor T. Polgar Fellowship Program
Kaili M. Gregory
Polgar Fellow
Environmental and Sustainability Sciences Cornell University Ithaca, NY 14853
Project Advisor:
Matthew Hare Department of Natural Resources Cornell University Ithaca, NY 14853
Gregory, M. K. and M. Hare. 2020. The Effect of Salinity on Eastern Oyster Reproduction in the Hudson River Estuary. Section IV:1-43 pp. In S.H. Fernald, D.J. Yozzo, and H. Andreyko (eds.), Final Reports of the Tibor T. Polgar Fellowship Program, 2018. Hudson River Foundation.
IV-1
ABSTRACT
Oysters have largely been missing from the Hudson River Estuary (HRE) since the early 1900s due to human activity, pollution and overharvesting. Water quality has been steadily improving in the estuary and coupled with the discovery of a wild remnant population, restoration is looking more feasible. Water quality factors such as dissolved oxygen, pH, and salinity can have effects on the survival and effectiveness of oysters.
This research focuses on the effect of salinity on the timing of gametogenesis in eastern oysters (Crassostrea virginica) between June and August 2018 in the Hudson River
Estuary (HRE). Gametogenesis was monitored in 2-year old oysters from experimental cage populations in the HRE. Adult oysters from a remnant population were transplanted to diverse salinities so growth rate and survivorship could be measured and compared.
Gametogenesis was evaluated using a histological gonad index, ranking the amount and maturity of gametes in the oyster gonad on a scale of 0-10. A GI value of 5 represents peak maturation, and a value of 6 represents active spawning. Based on literature, it was hypothesized that oyster gametogenesis would be delayed in lower salinity water relative to higher salinities in the Hudson River Estuary. The reproductive pattern was not a simple shift of the whole reproductive cycle, as predicted. A more dramatically altered reproductive phenology within the seasonal period studied was observed. The northern, low salinity sites started spawning earlier and spawned for longer through the reproductive season than the high salinity oysters. The results of this study demonstrate that adult oysters from the remnant wild population are flexible and tolerant of higher salinity conditions, and regardless of genetic strain, oysters change the timing of their reproductive output depending on the salinity where they live.
IV-2 TABLE OF CONTENTS
Abstract ...... IV-2
Table of Contents ...... IV-3
Lists of Figures and Tables ...... IV-4
Introduction ...... IV-5
Methods...... IV-11
Tappan Zee wild adult collection and transplant ...... IV-13
Monitoring of oysters and sampling for histology ...... IV-14
Histology preparation and Condition Index ...... IV-15
Histology slide analysis ...... IV-17
Statistical analyses ...... IV-18
Results ...... IV-19
Discussion ...... IV-27
Acknowledgments...... IV-33
References ...... IV-34
Appendix ...... IV-37
IV-3 LIST OF FIGURES AND TABLES
Figure 1 – Map of Hare Lab study sites ...... IV-12
Figure 2 – Shucked oyster anatomy...... IV-16
Figure 3 – Averaged weekly temperature data ...... IV-19
Figure 4 – Averaged weekly salinity data ...... IV-20
Figure 5 – Gonad index means and 95% confidence intervals - June ...... IV-22
Figure 6 – Gonad index means and 95% confidence intervals - July ...... IV-23
Figure 7 – Gonad index means and 95% confidence intervals - August ...... IV-24
Figure 8 –June GI vs. shell height ...... IV-25
Figure 9 – July GI vs. shell height ...... IV-25
Figure 10 – August GI vs. shell height ...... IV-25
Figure 11 – Gonad Index vs. Condition Index ...... IV-26
Figure 12 – Shell height over time (growth) for wild dredged TZ-HB adults ..... IV-27
Table 1 – Counts of oysters sampled for histology...... IV-14
Table 2 – Description of Gonad Index stages ...... IV-17
Table 3 – ANOVA results for effect on GI by all tested variables ...... IV-20
IV-4 INTRODUCTION
Crassostrea virginica, commonly known as the eastern oyster, was historically
abundant in the Hudson River Estuary. In the nineteenth and early twentieth century,
oysters were one of New York City’s top exports (Kurlansky 2006). Hudson River
oysters made it as far as Chicago, St. Louis, San Francisco, and even to Great Britain. In
1842, around $6 million worth of oysters were being sold in New York City. Adjusting
for inflation, that would be over $25 million in today’s economy. Eating oysters was
common and desired by people around the country (Kurlansky 2006). New York City
might be known today as the Big Apple, but back then the Big Oyster would have been
more appropriate.
As New York’s population grew, so did its pollution levels. Sewage and other types of waste were dumped into the estuary, harming the millions of oysters that lived on the bottom. Due to a synergistic mix of water pollution, eutrophication, disease and overharvesting, the once-abundant oyster beds near New York City were mostly dead by the beginning of the 20th century. Harvesting from the HRE beds was outlawed in 1920s
(Kurlansky 2006; MacKenzie 2007). Oyster populations are currently absent from most
of the Hudson River Estuary, but new data provides some hope for restoring this
keystone species to a more ecologically functional population despite the highly altered
conditions now existing in this urban estuary.
The water quality of the Hudson River Estuary has been steadily improving for
the past thirty years (Stinnette et al. 2018). Legislation such as the Clean Water Act of
1972 and the New York State Pure Waters Bond Act of 1965 prompted key initiatives to
clean up the Hudson. New sewage treatment plants have since been built along the
IV-5 estuary, and efforts by the New York State Department of Environmental Conservation and New York City Department of Environmental Protection have made positive strides towards cleaning up the HRE. The water is now generally safe to swim in, and more fish
species are entering the estuary (Stinnette et al. 2018). This long-term trend towards a
cleaner estuary system creates potential for restoration of a self-sustaining oyster population.
A successful restoration is characterized by a population large enough to be self- sustaining with measurable positive effects on ecosystem function. The eastern oyster is a keystone species, serving as a bioengineer that shapes its environment in ways that increase biodiversity and can indicate ecosystem health (Raj and Reson 2008). The population’s decline in the twentieth century left the estuary without this important biological and ecological keystone species.
The oyster’s reputation as a keystone species is based on several attributes.
Oysters are filter feeders, drawing water through their gills in order to obtain food. A single oyster can filter 50-55 gallons of water per day, removing particulates, detritus and dissolved pollutants (Raj and Reson 2008). Their effectiveness as filter feeders makes them reliable indicators of ecosystem health. Large populations of oysters in the
Chesapeake Bay have been shown to improve water quality in more shallow, mesohaline parts of the estuary (Coen et al. 2007). The water filtration process creates a positive feedback loop such that cleaner water promotes oyster survival and more diverse benthic communities. Additionally, the physical structure of an oyster bed provides habitat to numerous estuarine invertebrates and fish stocks that are economically important to humans (Peterson et al. 2003).
IV-6 A wild remnant population of Crassostrea virginica was discovered just south of
the Tappan Zee bridge in the Haverstraw Bay portion of the HRE (Figure 1) where
average salinity is low. Near the remnant population there has been consistently high
recruitment since observations began in 2012, but elsewhere in the HRE, only few and
scattered recruits are found in a typical year (McFarland and Hare, 2018). Until the
discovery of this wild remnant population, there was no known persistent reproductive
population in the HRE.
Discovery of the Tappan Zee-Haverstraw Bay (TZ-HB) remnant oyster
population gives greater hope to efforts focused on restoring oysters to the lower estuary
(closer to the ocean) because they need not start from nothing. As indicated by The
Nature Conservancy in their practitioner’s guide to restoration practice (zu Ermgassen
2016), wild oyster recruitment or transplants are a preferred method for restoration
compared to planting aquaculture or hatchery-produced ‘seed’ oysters (Brumbaugh et al.
2006). Hatchery propagation, even when done carefully from wild parents, typically results in genetic bottlenecks compared to wild populations (Appleyard and Ward 2006;
Hornick and Plough 2018). Oyster strains artificially selected for commercially valuable traits, such as fast growth, disease resistance, and attractive shells, are sometimes used in a restoration context (Baggett et al. 2014) but have unknown reproductive fitness in natural habitats. Restoration outcomes depend on fitness related performance over the entire life cycle. Thus, reproduction is important to measure and understand in order to evaluate the efficacy of different oyster seeding strategies in restored habitat.
Recruitment of wild-set oysters (newly-settled “spat”) has been monitored in the
TZ-HB region since 2008 and robust spat recruitment was observed in six of seven
IV-7 monitored years (Carthan and Levinton 2013; McFarland and Hare 2018; Starke 2010).
Additionally, ongoing research has shown that spat collected proximal to the wild TZ-
HB, when transplanted along the HRE salinity gradient and experimentally compared to
hatchery-produced oysters from mesohaline parents, show relative performance that
depends on location (McFarland and Hare, 2018). Preliminary findings suggest that the wild TZ-HB have superior growth and survival at the upper estuary low-salinity sites, but
equal or worse performance at harbor and Jamaica Bay (lower estuary) sites relative to an
aquaculture strain (McFarland and Hare 2018). These results suggest that the TZ-HB
oyster population may be locally adapted to their low salinity habitat as a result of long-
term isolation there. The research described here expands on these ongoing experiments
by quantifying and comparing one aspect of reproduction, the timing and extent of gonad
maturation.
The objective of this study is to understand how salinity and oyster strain (genetic
ancestry) affects reproductive timing for 2-year old oysters in the Hudson River Estuary.
Most restoration-related eastern oyster research has studied aspects of performance in the early parts of oyster life history (e.g., spat growth and survivorship), whereas reproduction has rarely been measured in natural eastern oyster populations. Studies have shown that salinities anywhere from 5-10 practical salinity units (psu) significantly slow- down or even inhibit the process of gametogenesis (Kennedy et al. 1996; Butler 1949); however, the same authors showed how tolerance to low salinity depends on the salinity at which broodstock were conditioned (at which gametogenesis occurred). It is challenging to study reproduction, especially in temperate waters where oysters need two
years to reach reproductive age. Histological measurement of gametogenesis stage is
IV-8 expensive, and fecundity is technically challenging to measure accurately (Kennedy et al.
1996; Loosanoff and Davis 1952; Mroch et al. 2012). The lack of studies on reproduction leaves great uncertainty about variation in critical fitness components: year of first reproduction, seasonal timing of spawn, age- and size-specific fecundity, and larval survivorship. Thus, sustainable oyster restoration in the HRE requires understanding of oyster response to a broad range of environmental conditions at every life stage: larva, juvenile, and adult.
Oyster reproduction is cyclic, separated into three stages generating the next generation of sessile juveniles: gametogenesis, spawning and external fertilization, and larval growth and settlement (Kennedy and Battle 1964). Gametes are synchronously released by adults so that zygotes are formed by external fertilization and then develop into early-stage larvae. The larvae spend 2-3 weeks in the water column, dispersing by tidal currents and advection. Available food resources (phytoplankton) will determine the duration of the larval period before settlement competency is developmentally accomplished. The eventual settlement location of oyster larvae is hard to predict because environmental factors such as water currents and predators vary among sites. After oyster
larvae become competent to settle, they ‘swim’ to the bottom using ciliary propulsion,
where they find a hard substrate and metamorphose into sessile spat (Kennedy et
al.1996).
Eastern oysters are protandrous hermaphrodites, maturing first as males and later
becoming females (Kennedy et al. 1996). In temperate waters C. virginica spends most of
the winter as an undifferentiated sex (Kennedy et al.1996). Sex differentiation, spawning,
and reproduction follow a seasonal pattern. Gametogenesis builds gonad tissue through
IV-9 the Spring, and at a temperature between 20-30°C, the oysters begin to release gametes
into the water column (Kennedy et al. 1996; Barber et al. 1991). The oysters spawn
synchronously, so when one releases its gametes, nearby oysters are triggered to release
their own. (Mann et al. 2014; Loosanoff and Davis 1952). After the oysters have
spawned, they resorb their follicles and re-enter the sexually undifferentiated stage in the fall. At this time follicles become very small and separated from each other by lots of connective tissue. At this stage, no differentiated sex cells are visible; however, some slight gametogenic activities might still be occurring. As it gets colder, the oysters enter a torpor stage in which gonads are dormant until the next Spring (Kennedy et al.1996;
Loosanoff 1942).
Because of its importance for aquaculture, the thermal induction of oyster spawning is well studied. This study will examine the second-order effect of salinity in a natural system (Volety 2008; Loosanoff and Davis 1952). Studies in a natural system are vulnerable to potentially confounding variables such as nutrient levels and pollution that can make it challenging to identify specific environmental triggers or thresholds. The effects of salinity, in particular, are important to study in the HRE because of the potential utility of transplanting oysters from the wild remnant low-salinity TZ-HB population to southern, moderate to high salinity portions of the estuary for restoration.
Oysters are tolerant of a broad range of salinities (~ 5 – 40 psu), so they typically occur across much of the salinity gradient within estuaries. Results from the HRE may provide observations that are more generally true, at least in temperate estuaries.
There is agreement among previous research that lower salinities limit or even inhibit reproduction in oysters. A 1949 study found that oyster reproduction is entirely inhibited
IV-10 under 6 psu (Butler 1949).) A study by Volety found that oysters at a salinity below 14 psu demonstrated poor spat recruitment, excessive valve closure and slower growth. The smaller oysters were less fecund, producing significantly fewer eggs than their larger, higher salinity (14-28 psu) counterparts.
Relative to the limits suggested by these early studies, it seems contradictory that the wild TZ-HB population currently exists at salinities ranging from 0-12 psu, depending on season. The TZ-HB population’s existence suggests that C. virginica reproduction might be more phenotypically plastic than originally thought, or that the
TZ-HB population may be uniquely adapted to the low salinity environment.
Based on this literature and the observation that peak spat recruitment is often in
September for the TZ-HB wild population (McFarland and Hare 2018), the hypothesis tested in this study was that oyster gametogenesis is delayed in lower salinity water relative to higher salinities in the Hudson River Estuary. The delay might also be related to the timing of Spring water warming, which is delayed in the upper estuary relative to the lower estuary. This study evaluates gonad maturation relative to three main factors: sex, location (i.e., salinity), month, and oyster strain in the HRE.
METHODS
As stated above, temperate latitude eastern oysters require two years of growth before they become reproductive. Therefore, the objective of this project was to study the eastern oyster cohorts out-planted as one-month old juveniles in August 2016 by the Hare
Lab. Over the past two years, the Hare Lab has monitored survivorship and growth for three oyster strains (3 experimental cohorts with different parentage) maintained in
IV-11 experimental cages hung from docks. These 2016 cohorts are independent replicates of
initial experiments run from 2015 to 2017 at a subset of the sites (McFarland and Hare
2018).
Four outplant sites were used for
sampling 2 year old experimental oysters in
this study (Figure 1). They included “river”
sites at Hastings-on-the-Hudson (HH) and
Yonkers Science Barge (SB), a “harbor” site
located at Red Hook, Brooklyn (RED), and a
Jamaica Bay site at the Sebago Canoe Club in
Paerdegat Basin (PGB). Temperature and
salinity were recorded every two hours near
each of these sites using YSI 600 sondes
deployed by the Hare Lab or using a nearby
Hudson River Environmental Conditions Figure 1. Map of Hare Lab oyster sampling study Operating System (HRECOS) site. Both the sites along the HRE salinity gradient. sondes and the experimental cages were hung
1m below a dock or were on a rocky bottom in the low intertidal zone (Hastings). Data from Piermont Pier, which is near SB and HH was obtained from the HRECOS (http://hudson.dl.stevens-
tech.edu/hrecos/d/index.shtml). Sonde data from Kingsborough Community College
(KCC) in Jamaica Bay was used to represent environmental conditions at the nearby
Jamaica Bay (PGB) site.
IV-12 There are three different cohorts being compared in this study: wild, hatchery and
aquaculture, all the same age (within 1 month). The wild oyster sample was obtained as
August or September wild-set recruits on bivalve shell in the vicinity of the wild TZ-HB
remnant population. The hatchery cohort was produced by strip-spawning wild Martha’s
Vineyard oysters from a moderate salinity lagoon (similar to methods in McFarland and
Hare 2018). The larvae were then cultured by the Martha’s Vineyard Shellfish Group and
sent to the Cornell Cooperative Extension hatchery in Southold, New York to set on shell
in an upwelling system for three weeks during August 2016 before out planting to the
HRE cages. Seed oysters from a domesticated aquaculture strain also were acquired in
August 2016. Aquaculture seed and hatchery-produced oysters were kept in small-mesh bags until large enough to be contained in poly mesh netting bags.
Tappan Zee Wild Adult Collection and Transplant
Wild TZ-HB adults were desired in order to transplant them to different salinities and measure resulting variation in their gonad maturation. The oysters were dredged in cooperation with Dr. Tiffany Medley, using Monmouth University’s R/V Seahawk. A total of 276 live, wild adult TZ-HB oysters were dredged on June 15, 2018 for this study.
The oysters were stored in a cooler with water from the dredging location overnight. On
June 16, the oysters were randomly distributed in groups of 50-65 to the following sites:
HH, SB, LM, RED, and PGB (Fig 1). Unfortunately, logistical constraints prevented collections before June 15. It was decided that June 15 was too far into the reproductive maturation season to cleanly interpret reproduction after transplant, so the TZ-HB wild
IV-13 adults were monitored for variation in growth rate and survivorship and held for
experiments in 2019.
Monitoring of Oysters and Sampling for Histology
Samples for histology were collected three times during the summer of 2018: June
15-21, July 8-13, and August 8-13. At each site visit, growth and survivorship were measured for each cohort. Shell height was measured to the nearest millimeter using
Mitutoyo Calipers (Mitutoyo America Co., Aurora, IL, USA). Fouling organisms were cleaned off cages at each visit, but not from oysters unless they interfered with measurements. For histology, a maximum of 20 oysters were collected from each cohort at each site (Table 1). The number collected was limited by cohort abundance and budget.
Planned sample size variation prioritized July and August samples, and poor survivorship
at some sites prevented complete sampling.
Table 1. Counts of oysters sampled for histology from each cohort at each site during each month. Site HH SB RED PGB
Jun Jul Aug Jun Jul Aug Jun Jul Aug Jun Jul Aug
AQ 13 20 20 13 20 20 11 18 20 13 19 20
Hatchery 13 12 12 13 20 20 0 0 0 13 18 20
Wild 13 20 19 13 20 20 0 8 6 13 20 20
IV-14 Histology Preparation and Condition Index
Oysters were transported in coolers with ambient HRE water to the Fort Totten
Urban Field Station lab (US Forest Service) where dissection and preservation was done.
Lab work was performed as close to sampling date and time as possible in order to ensure that the histology results reflected the oyster’s response to their respective site’s environment, not the environment of the cooler used to transport them. Most processing was same-day, or at most the morning after collection.
The oysters were cleaned with a wire brush in order to remove any barnacles, dirt or plant tissue. Shell height was measured (again, repeating field measures) and then the oyster was shucked carefully so as to not mangle the oyster’s tissue. Using a scalpel, the oyster was removed from the shell, taking care to not damage the soft tissue. For each oyster, a new razor blade was used to make a single clean, diagonal cut across the body of the oyster, and then another cut along the same axis, but 4mm from the initial cut
(Figure 2). The oyster slice was placed in a labeled plastic cassette and preserved in
Davidson’s Solution for 7 days (Volety 2008). As much as possible, the position of the cross section was standardized among all individuals. After fixation and a 70% ethanol wash, the samples were sent to the Cornell University Animal Health Diagnostic Center for histology slide preparation with hematoxylin and eosin staining.
In bivalves a useful indicator of overall physiological condition is the condition index, a measure of the ratio of dry tissue weight to dry shell weight. Excess energy that is not spent on reproduction or daily metabolism is invested into the oyster’s biomass.
IV-15
Figure 2. Shucked oyster anatomy showing orientation (parallel lines) of the section taken for histology preparation (Howard et. al, 2004). Whereas shell growth is cumulative with age, soft body tissue shows seasonal mass
variation on top of cumulative growth; spring-summer gonad maturation often generates a peak in soft tissue mass and in condition index (Ruiz et al. 1992). Environmental stressors can cause oysters to invest less in growth, thus affecting condition index (Volety
2008).
Condition index was measured using all soft oyster tissue except the histology slice. Thus, CI in this study may have inflated variance due to variation in the proportional mass of histology slices, and it is not comparable to CI in other studies.
Oyster soft tissue was placed in a labeled, pre-weighed aluminum dish and along with both shells was placed in a drying oven at 70 °C for approximately 36 hours. Condition index was determined from the dry weights, using the following formula (Lucas and
Beninger 1985; McFarland and Hare 2018).
dry tissue weight (g) CI = × 100 dry shell weight (g)
IV-16 Histology Slide Analysis
The histology slides were examined with a compound microscope under 10X and
40X magnification to determine sex and gametogenic stage. Gonad index (GI), a method for representing gametogenic stage, was scored using the criteria described in Table 2.
Pictures of each gonad stage for each gender were taken using Leica LAS Microscope
Software. The pictures can be found in the Appendix.
Table 2. Description of Gonad Index stages (derived from Volety 2008). Stage Description 0 Neuter or indeterminate sex despite good histology preparation; no presence of follicle or connective tissue 1 Determinate sex indicating beginnings of gametogenesis; no mature gametes visible 2 Females: no more than one-third mature eggs relative to developing eggs Males: fringe follicles starting to accumulate mature gametes; low density of sperm 3 Females: no more than one-half mature eggs relative to developing eggs Males: visible connective tissue; visible sperm tails in center of some follicles 4 Females: mostly mature polygonal eggs and distended follicles with some developing eggs still visible Males: small amount of visible connective tissue; compact follicles; sperm tails visible in most follicles 5 Females: only mature polygonal eggs; no empty spaces from spawning gametes Males: sperm have visible tails; uniformly high sperm density in packed follicles; no visible connective tissue 6 Active spawning is occurring; follicle structure is disrupted Females: general rounding of eggs; a few empty spaces from released eggs Males: reduction in sperm density; sperm tails less visible 7 Follicles one-half depleted of gametes Males: sperm area reduced to one-half of gonadal area 8 Follicles two-thirds depleted of mature gametes; further reduction of gonadal and connective tissue area 9 Only residual gametes remain; determinate sex; further reduction of gonadal and connective tissue area 10 Gonads devoid of most or all gametes; connective tissue is still visible but very minimal; sex not always determinate Females: one or two visible eggs remain Males: connective tissue has a few remaining sperm to allow for sex identification
IV-17
Statistical Analyses
A random linear model was created based on the gonad index results to predict multiple interaction effects of sex, location, strain, and month on GI. For this model, the
GI values from 6-10 were relabeled to 5-1 (6 = 5, 7 = 4, 8 = 3, 9 = 2, 10 = 1) to allow for averages that reflect the true gonad condition. For example, two oysters with gonad indexes of 2 and 9, respectively, would have a meaningless “unfolded” average GI of 5.5.
Neither of the oysters are at peak reproductive spawning, yet that is what the “unfolded” average would suggest. For the folded index with 9=2, the “folded” average in this example would be 2, reflecting the oyster’s true relation to peak reproduction (Volety
2008). A dummy variable [0,1] was assigned to the GI values before “folding”, called ‘GI trend’ to represent the gametogenesis and spawning GI distributions. GI values 1-5 were assigned GI trend=0 to represent oysters that are still actively growing gonads, or gametogenesis. GI values 6-10 were assigned GI trend=1 to represent oysters that are spawning out or resorbing their gametes. GI values of 0 were ignored for the GI trend variable because it was impossible to confirm if an oyster with a GI of 0 had not undergone gametogenesis or had completely resorbed gametes. A one-way Analysis of
Variance (ANOVA) was performed on the model that included the effects of sex, location, strain, month, and GI trend on GI.
Results were deemed significant at P < 0.05. Predicted means and confidence intervals were generated using the random linear model and plotted for select combinations of variables. A Spearman’s test of correlation was used to determine the
IV-18 relationship between GI and shell height. Statistical analyses were conducted using
RStudio (RStudio Team 2016).
RESULTS
Temperatures ranged from 16-28 °C in the summer months. Surprisingly, HH had the highest temperatures, even though it was the northernmost site. A study by Loosanof identified 20 °C as the temperature at which oysters spawn, which sets up the prediction that when oysters cross this threshold, they will begin to spawn (Loosanof and Davis
1952). HH is the first site to cross the threshold, followed by KCC and RED. SB temperature data was not available in the earlier part of the study period. Environmental data was not available for SB until the beginning of July.
Figure 3. Averaged weekly temperature data with standard error bars. The dashed horizontal line represents the temperature threshold for spawning in temperate oysters (Loosanof and Davis 1952)
IV-19 Predictably, salinity was lowest at the two southernmost sites: RED and KCC.
Their salinities ranged from 16-27 psu. HH had the lowest salinity, followed by RED.
The salinity data matches the known salinity gradient in the estuary (Figure 1).
Figure 4. Averaged weekly salinity data with standard error bars.
Table 3. ANOVA results for effect on GI by all tested variables. ** denotates a significant P value. Variable P values Sex 0.2227975 Strain 0.0842607 Month 7.725e-10 ** Location 0.9357032 GI trend 3.508e-05 ** Strain*GI trend 0.9443797 Month* GI trend 5.245e-07 ** Location*GI trend 0.0464173 ** Month*Location 4.044e-09 ** Month*Location* GI trend 0.0008605 **
IV-20
In the one-way Type III ANOVA test, strain did not have a significant effect, even in interaction with GI trend. As expected, due to temperate latitude seasonality, GI varied significantly between months, and there was also an interaction between month and GI trend. Location had a significant effect on the GI means only in the context of an interaction with GI trend or GI trend and month.
Graphs of means and 95% confidence intervals were created in order to observe
GI distribution differences. If the confidence intervals did not overlap, it was determined that the difference between the compared groups was significant. The ANOVA results
(Table 3) reveal that there is a significant difference between the GI trend values 0 and 1, or gametogenesis and spawning GI patterns, respectively. Therefore, for visualization, the
GI data was ‘unfolded’ by separating gametogenesis (0-5) and spawning (6-10) means in
Figures 5, 6 and 7. The ANOVA test also demonstrated that strain had no significant effect, so it was not included in the confidence intervals.
The June gametogenesis data demonstrates a significant difference in gonad maturation between the highest salinity site (PGB) and the two lowest salinity sites (HH and SB). The lower salinity sites, which are closer to the location of the wild remnant population, had a higher gametogenesis GI, indicating that the oysters were farther along in the gametogenesis than the oysters at the lower salinity sites. The site with most advanced reproduction was Science Barge (SB), with almost half the sample in the spawning stage. The other sites had very few individuals in the spawning stage. The RED data for this and other months show large confidence intervals due to especially high variance in GI values observed in oysters at this location.
IV-21
Figure 5. Gonad index means and 95% confidence intervals plotted for each site for the month of June. Labels of the number of gametogenesis and spawning oysters for each site are to the right of their corresponding interval and mean. A vertical dashed line separates the two sides of the ‘folded’ GI, approximately corresponding to gametogenesis and spawning stages.
Red Hook was the outlier in July, even after accounting for large confidence
intervals around the GI means. In the gametogenesis GI stages, there was no difference
between the lower and higher salinity sites. Compared the June, the oyster sample at PGB
is much more advanced in terms of gametogenesis GI. PGB in Jamaica Bay had a higher
number of spawning oysters than river sites and a significantly higher GI spawning mean
(GI = 7.0) than HH. The spawning GI average for HH and SB were approximately the same in June and July. Overall, there are more oysters in the spawning stage in July across all sites.
IV-22
Figure 6. Gonad index means and 95% confidence intervals plotted for each site for the month of July. Labels of the number of gametogenesis and spawning oysters for each site are to the right of their corresponding interval and mean. A vertical dashed line separates the two sides of the ‘folded’ GI, approximately corresponding to gametogenesis and spawning stages.
In August, the river sites (HH and SB) continued to show evidence of spawning in less than half the sample, whereas more than 2/3 of the oysters at RED and PGB had spawned. There is no difference between sites in the gametogenesis stage. As seen in
July, spawning PGB oysters were significantly more advanced than at river sites with an average GI (8.64) closer to 10 than HH (7.42) and SB (7.69).
IV-23
Figure 7. Gonad index means and 95% confidence intervals plotted for each site for the month of August. Labels of the number of gametogenesis and spawning oysters for each site are to the right of their corresponding interval and mean. A vertical dashed line separates the two sides of the ‘folded’ GI, approximately corresponding to gametogenesis and spawning stages.
To explore whether reproductive timing might be influenced by oyster size, shell height was plotted against gonad index and separated by month. A Spearman’s test of correlation was performed for each relationship with GI for June, July and August. In
June and August there was no significant monotonic relationship between oyster size and gonad maturation stage (Figures 9,11). For July, the P value was 0.025 and the rho value was 0.160 for a positive correlation (Figure 10).
IV-24
Figure 8. June GI vs. shell height. Figure 9. July GI vs. shell height.
Figure 10. August GI vs. shell height.
IV-25 To observe the relationship between oyster condition and gonad maturation, GI was plotted against condition index (Figure 11). Shucking procedures occasionally resulted in broken shell that was unrecoverable for condition index measurement. As expected, Figure 11 shows that condition index peaks when the gonad index is around 5 or 6, representing the ripest reproductive stages with the greatest soft tissue mass.
Figure 11. Gonad Index vs. Condition Index.
Growth rate and survivorship of dredged TZ-HB oysters was measured in October as well as June - August. The wild TZ-HB oysters continued to grow once they had been transplanted (Figure 13). At RED, there was a 45% mortality rate between August and
October. Mortality was very low at all other transplant sites (< 7.4%).
IV-26
Figure 12. Shell height over time (growth) for wild dredged TZ-HB
adults transplanted to five sites in the Hudson River
Estuary.
DISCUSSION
The results of the random linear model and confidence intervals do not lend support to the simple hypothesis that gametogenesis is delayed in lower salinities along the Hudson River Estuary; however, there is an observable difference between the reproductive timing of oysters at different locations along the salinity gradient. In June, few oysters were spawning and the locations with lower salinity had more oysters closer to peak spawning (GI = 5) on average than the oysters at higher salinity locations. Later in the reproduction season, in July and August, the post-spawn oysters at high salinity sites had significantly higher GI values (more depleted and reabsorbed gonads) relative to those at low salinity sites. Together, these observations suggest that oysters at higher salinities had a delayed, but more rapid spawning period relative to oysters at low salinity. This was also apparent in August by the proportion of oysters that had not yet
IV-27 spawned. River sites at low salinity had more than half pre-spawned individuals whereas the harbor (RED) and Jamaica Bay (PGB) sites were mostly spawned-out. In other words, the northern low salinity sites seemed to start spawning earlier and extend reproduction later, compared with the harbor and Jamaica Bay sites. These findings for
River sites are consistent with the September-October spat set reported for the TZ-HB area (McFarland and Hare 2018).
The four sites were arranged along a salinity gradient, but spatially positioned such that the two northern sites (HH and SB) had similar and significantly lower salinity values (5 – 15 psu) than the southern sites (18 – 26 psu; Figure 4). There was some temperature variation among sites, but at the first collection in June the range of temperatures was 18 – 21 °C across sites, close to or above the threshold temperatures that have been reported from temperate waters. It is possible that during April and May, when both average temperature and salinity gradients are steeper than in June, early gametogenesis may have been more advanced at the mesohaline sites, which crossed the temperature threshold earlier than the southern sites (Figure 3; Loosanof and Davis
1952).
Overall, the observed temporal pattern of reproduction documented over the summer months in the Hudson River Estuary supports the expectation that oysters in temperate regions will spawn with a major peak during the summer months and enter a non-reproductive stage in the winter months (Cox and Mann 1992; Loosanof and Davis
1952). The sampling constraints in this study made it impossible to evaluate whether some oysters redeveloped gametes for a Fall spawn. This is one of the first reports that shows the degree of temporal discreetness of a seasonal peak in reproduction to be
IV-28 dependent on salinity. This finding implies that hatcheries will be more successful getting oysters to condition uniformly if done at moderate to high salinity.
In the ANOVA test, Location did not show significance on its own, but this is likely because not all locations were different from each other. Some of the locations had very similar GI data (HH and SB, RED and PGB). The ANOVA does not test for differences between individual locations, which is why means and confidence intervals were calculated. When Month and GI trend were taken into account with Location, represented by Month*Location* GI trend, and there was significance in the linear model.
Data from September and October would have been valuable to observe how long it takes the oysters to completely spawn and enter their torpor period. A longer sampling period would also be interesting to see if any locations supported a Fall resurgence in gametogenesis and spawning, as reported in the mid-Atlantic and northern Gulf of
Mexico (Southworth and Mann 2004; Hayes and Menzel 1981).
There is potential for measurement error in this study related to the GI scoring.
Even with clearly defined categories, there is some subjectivity in scoring intermediate specimens. GI values of 0 and 10 do not contain any gametes and can be confused with each other. GI= 10 is slightly different in that some follicle tissue remains. In addition to multiple GI evaluation trials, the oysters with GI values of 0 and 10 were closely examined to determine the correct scoring. Incorrect scoring between the 0 and 10 is possible, which would cause the average GI values and confidence intervals to be skewed. GI values of 0 were not included in the ANOVA test or confidence intervals, but values of 10 were because of the remaining follicle tissue. Small and uneven sample sizes
IV-29 limit the ability to precisely quantify and compare gametogenesis state. Most of the broad
confidence limits in Figures 4, 5 and 6 are due to especially small sample sizes.
The results of this study offer a unique examination of strain, month and location
effects on gametogenesis timing for C. virginica along a temperate estuarine salinity
gradient. It is not clear how to reconcile results from this study with the findings of Butler
(1949), who found that gametogenesis was inhibited in 90% of his study population when
salinity was below 6 psu. At the northern most site included in this study, HH, average
weekly salinity reached briefly down to 5 psu or slightly below. Previous studies have
found that the TZ-HB region regularly experiences Spring salinities below 5 psu,
sometimes for multiple consecutive weeks during the spring snow melt (McFarland and
Hare 2018). Thus, oysters at the low salinity River sites in this study may have
experienced inhibition of gametogenesis earlier in the year before sampling began. If that
is true, it implies that oysters can phenologically compensate for environmental
conditions that delay gametogenesis (cold temperature and/or low salinity).
The aquaculture, hatchery and wild strains were all able to spawn gametes and
showed similar gametogenic phenology. This result suggests that the variation in
reproductive timing observed in this study is not genetic, but a result of acclimation and phenotypic plasticity. Therefore, with respect to reproduction, transplanted oysters can be expected to adjust the timing of their gametogenesis to acclimate to new salinity and temperature conditions. The experimental oysters studied developed from an age of ~4 weeks at the site from which they were sampled. Uncertainty remains about how long acclimation would take after a transplant and whether there are key developmental periods that facilitate acclimation. These findings are based on a comparison of three
IV-30 temperate oyster strains, and therefore do not contradict the previous conclusion, based on common garden studies, that genetically-based physiological races occur between more latitudinally disparate source populations (Barber et al. 1991).
The significant but weak monotonic relationship between gonad index and shell height suggests that larger oysters progress through the reproductive cycle sooner than smaller oysters. The power to detect a correlation was highest in July, with a weak monotonic relationship was observed in (rho = 0.160). This positive correlation implies that larger same-age oysters are slightly more advanced in gametogenesis (and resorption) than smaller oysters at a given time, on average. Mostly studies have measured the positive correlation between fecundity (egg count) and shell height (Mann et al. 2014; Mroch et al. 2012), making these findings unique.
Condition index is a ratio of tissue weight to shell weight, varying with seasonal changes in physiological status, especially reproduction (Volety 2008). When oysters begin to produce gametes, their soft tissue weight increases, thus increasing the value of the condition index. When plotted against GI, condition index increased as GI increased from 1-5. Interestingly, condition index did not increase much between GI 5 and 6, even though by definition, oysters only began to release gametes at GI 6 (Fig. 11). This implies that gonad mass is not increased during the final stage of gametogenesis.
Although GI was not measured for the transplanted TZ-HB wilds, growth and survivorship provide valuable information about how these wild adult oysters performed in salinities different than where they settled and matured. There does not appear to be any evidence for a dredging or transplant affect depressing growth, even when transplanted to higher salinity at PGB. Growth and survivorship were not significantly
IV-31 different among locations during the summer months, but examination in October
revealed performance differences. The oysters at RED had significantly lower growth and
survivorship in October. Over 45% of oysters died at RED whereas most oysters survived
at the other 4 sites. The pattern of poor survivorship at RED for TZ-HB wild adult oysters
also was observed in other cohorts at this site (McFarland and Hare, 2018). Moderate
salinity at RED is ideal for eastern oysters, but there are many factors that could account
for this trend such as harbor pollution, low nutrient availability, high sediment load and
disease. More in-depth experimentation and environmental monitoring at this site would
be necessary to pinpoint the cause of poor survivorship.
Besides mortality at RED, the growth and survivorship of TZ-HB oysters at the
other transplant sites demonstrates potential for TZ-HB oysters to be a source for
successful transplantations. Although wild local oysters are preferred for restoration
efforts to capitalize on any local adaptations, comparisons among strains here indicate
that gametogenic phenology may be suitable in any temperate oyster strain, whether wild
or hatchery produced, if allowed a suitable period for acclimation. Further study would
need to be done to compare other fitness components and to examine the population
genetic effects of using hatchery-produced or domesticated oysters in restoration. Firm
conclusions will require a more complete evaluation of fitness for transplanted oysters
over the entire life cycle. These experimental transplants have created the opportunity for
further studies to determine how transplanted adults from a low-salinity area survive and grow along the HRE salinity gradient, and by spawning them in the future it will be possible to test relative performance of their larvae. This study is only the beginning of
IV-32 the necessary amount of information to inform conservation and repopulation efforts in the Hudson River Estuary.
ACKNOWLEDGEMENTS
We would first like to thank the Hudson River Foundation for the opportunity and funding of the Tibor T. Polgar fellowship. Katie McFarland, Sarah Gisler, and Tiffany
Medley supported this research by offering advice, help with field work, and data analysis consults. We are grateful to Captain Jim Nickels for leading a successful oyster dredging trip. Thanks also to the Cornell Statistical Consulting Unit for advice on the data analysis and graphical representation for this research. We thank the U.S. Forest
Service Fort Totten Urban Field Station for providing housing and a lab for this field work. Finally, this work would not be possible without the generous site hosts that keep
Hare Lab experimental oyster cages safe.
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IV-36 APPENDIX
Photographs of oyster gonads taken on Leica LAS Microscope Software. “1F 25x” is a female oyster with a GI of 1 taken at 25x.
1F 25x 1F 100x
1M 25x 1M 100x
2F 16x 2F 80x
IV-37
2M 12.5x 2M 50x
3F 10x 3F 90x
3M 16x 3M 40x
IV-38
4F 10x 4F 63x
4M 12.5x 4M 40x
5F 10x 5F 90x
IV-39
5M 12.5x 5M 25x
6F 12.5x 6F 80x
6M 10x 6M 40x
IV-40
7F 12x 7F 80x
7M 16x 7M 50x
8F 12.5x 8F 80x
IV-41
8M 9x 8M 32x
9F 12.5x 9F 100x
9M 8x 9M 32x
IV-42
Lower resolution 10F photo unavailable 10F 63x
10M 12.5x 10M 55x
0 25x 0 80x
IV-43