University of Central Florida STARS

Electronic Theses and Dissertations, 2004-2019

2005

Native And Invasive Competitors Of The Eastern Oyster Crassostrea Virginica In Mosquito Lagoon, Florida

Michelle Boudreaux University of Central Florida

Part of the Biology Commons Find similar works at: https://stars.library.ucf.edu/etd University of Central Florida Libraries http://library.ucf.edu

This Masters Thesis (Open Access) is brought to you for free and open access by STARS. It has been accepted for inclusion in Electronic Theses and Dissertations, 2004-2019 by an authorized administrator of STARS. For more information, please contact [email protected].

STARS Citation Boudreaux, Michelle, "Native And Invasive Competitors Of The Eastern Oyster Crassostrea Virginica In Mosquito Lagoon, Florida" (2005). Electronic Theses and Dissertations, 2004-2019. 532. https://stars.library.ucf.edu/etd/532 NATIVE AND INVASIVE COMPETITORS OF THE EASTERN OYSTER CRASSOSTREA VIRGINICA IN MOSQUITO LAGOON, FLORIDA

by

MICHELLE LEAH BOUDREAUX B.S. University of South Florida, 2003

A thesis submitted in partial fulfillment of the requirements for the degree of Master of Science in the Department of Biology in the College of Arts and Sciences at the University of Central Florida Orlando, Florida

Fall Term 2005 ABSTRACT

Populations of Crassostrea virginica within Mosquito Lagoon, Florida have recently

undergone significant die-offs, which are a subject of major concern. Restoration efforts

within Mosquito Lagoon are focusing on reconstructing the three-dimensional reef habitats.

Before effective protocols can be established, however, important questions about the

sources of juvenile and adult oyster mortality must be answered. Potential causes of

Crassostrea virginica mortality in the Indian River Lagoon system include sediment loads,

competition, predation, and disease. My research focused on the interactions between

oysters and the competitors that may affect the settlement, growth, and survival of Crassostrea

virginica. The four objectives of my thesis research were to: 1) identify potential oyster

competitors in Mosquito Lagoon, 2) determine if the sessile species recruiting to oyster shells

have changed over time, 3) determine how the dominant competitors, barnacles, affect

oyster settlement, growth and survival, and 4) determine if oyster or barnacle larvae are

better able to settle in increased sediment and flow conditions that are associated with high

levels of recreational boating.

Lift nets were deployed within Mosquito Lagoon to determine available competing

species. I collected species inventory data at six sites to determine the sessile invertebrate

species (competitors) present on oyster reefs. Nets were deployed intertidally, just above

mean low water, on living oyster reefs. One and a half liters of live and dead oysters were

placed within the nets upon deployment. The nets were picked up monthly and surveyed for all fauna. Upon retrieval, all oysters within each net were brought back to the lab where all sessile organisms were immediately identified and returned to the lagoon. This survey began June 2004 and continued for one year.

ii Shells from historic shell middens (up to 15,000 years old) were examined to determine if the sessile species settling on oyster reefs have changed over time. Similar species were found on both shells of historic and extant reefs. One notable exception was the appearance of Balanus amphitrite, an invasive barnacle, on the extant reefs. Balanus amphitrite is thought to have invaded Mosquito Lagoon approximately 100 years ago. This has resulted in a five fold increase in barnacle abundance per oyster shell.

Balanus spp. were identified as important potential competitors and thus my research focused on spatial competition between C. virginica and native versus invasive barnacles of the area. Over 300 barnacles, including a native species, Balanus eburneus, and an invasive,

Balanus amphitrite, have been counted on a single oyster shell. To determine how Balanus spp. affected settlement, growth, and survivorship of C. virginica, laboratory and field experiments were conducted in which densities of Balanus amphitrite and Balanus eburneus were manipulated. Density treatments included: no barnacles (control), low, medium, and high coverage of barnacles. Laboratory settlement trials with cultured oyster larvae were run in still water and flow (recirculating flume) using all barnacle density treatments.

Additionally, all treatments with 7-day oyster spat were deployed in the field to follow oyster spat growth and survivorship. Settlement was counted by microscopy, and growth and survivorship were measured every 3 days for 4 weeks. Settlement of oysters was affected by barnacle presence only in flowing water. Still water trials showed no oyster preference related to any barnacle density or species. The presence of barnacles affected the growth and survivorship of oyster spat. However, there were no species specific differences.

Studies suggest that recreational boating activities, especially boat wakes that cause sediment resuspension, may decrease recruitment and this may then provide an advantage to sessile competitors less affected by flow and sediment loads. To address these issues,

iii replicated laboratory trials were run in a laboratory flume to quantify the effects of water motion (0, 5, 10 cm/s) and sediment loads (0, 8, 16 g/ml) on oyster recruitment and the recruitment of an important, relatively new competitor in the system, the barnacle Balanus amphitrite. If B. amphitrite settles in a wider variety of flow rates and sediment conditions, it may have a competitive advantage over the native oyster in this space-limited habitat. I found that high flow and sediment loads reduced larval settlement of C. virginica.

Alternatively, settlement of cyprids of B. amphitrite did not differ among treatments. Thus, continuous boat traffic during settlement times should favor recruitment of the invasive barnacle Balanus amphitrite over the native oyster Crassostrea virginica.

Determination of the competitive interactions of Crassostrea virginica in Mosquito

Lagoon gives us important insights into the ecological conditions necessary for re- establishment of these oyster populations. Crassostrea virginica in Mosquito Lagoon was significantly impacted by barnacles; settlement, growth, and survivorship were all reduced by

Balanus spp. This information will help resource managers in planning restoration techniques to minimize oyster and barnacle competitive interactions and increase Crassostrea virgininca success.

iv ACKNOWLEDGMENTS

Funding was provided by the University of Central Florida Biology Department, including a

Research Enhancement Grant, Sigma Xi Grant in Aid of Research, and Florida Sea Grant

(Linda Walters). I thank the University of Central Florida and Canaveral National Seashore for their support in this project. I would like to thank my thesis advisor Dr. Linda Walters as well as my committee members, Dr. Jack Stout and Dr. Mark Luckenbach. I greatly appreciate all those who helped with data collection: J.L. Stiner, A. Barber, M. Black, M.

Donnelly, N. Gillis, C. Glardon, J. Ledgard, A. McLellan, J. Sacks, P. Sacks, A. Simpson, E.

Stiner, J.C. Stiner, and J.K. Stiner. I thank the following for other assistance making this project possible: Southeastern Archeological Center, Dr. Benson, Dr. Lee, Dr. Rittschof, B.

Orihuela, Dr. Arnold, Dr. Scarpa, K. Kurkowski, Dr. Tolley, Dr. Volety and Dr. Quintana.

Finally, I would like to thank my family and friends for their endless support through this project.

v

TABLE OF CONTENTS LIST OF FIGURES...... vii LIST OF TABLES ...... ix GENERAL INTRODUCTION ...... 1 Indian River Lagoon System ...... 1 Importance of Crassostrea virginica...... 3 Biology of Crassostrea virginica...... 6 Crassostrea virginica in Mosquito Lagoon ...... 19 CHAPTER ONE: BIODIVERSITY OF SESSILE SPECIES ON OYSTER REEFS...... 20 Introduction...... 20 Methods: Lift Nets (Present Day Species) ...... 21 Results...... 24 Methods: Middens (Historical Analysis of Organisms Attached to Oysters)...... 26 Results...... 27 Discussion...... 28 CHAPTER TWO: EFFECT OF BARNACLES ON OYSTER SETTLEMENT, GROWTH AND SURVIORSHIP...... 31 Introduction...... 31 Methods...... 34 Results...... 37 Discussion...... 38 CHAPTER THREE: EFFECT OF FLOW AND SEDIMENT ON THE SETTLEMENT OF OYSTER AND BARNACLE LARVAE...... 40 Introduction...... 40 Methods...... 42 Results...... 46 Discussion...... 46 APPENDIX A- FIGURES...... 49 APPENDIX B- TABLES...... 71 REFERENCES...... 87

vi LIST OF FIGURES

Figure 1- Indian River Lagoon system (Smithsonian 2001) ...... 50 Figure 2- Oyster reefs...... 51 Figure 3- Internal diagram of Crassostrea virginica (Ashbaugh 1951)...... 52 Figure 4- Life-cycle of Crassostrea virginica (Wallace 2001)...... 53 Figure 5- Lift nets study sites in Mosquito Lagoon, Florida...... 54 Figure 6- Submerged lift net (1m2) ...... 55 Figure 7- Lift net retrieval...... 55 Figure 8- Mean richness (total number of species) ± S.E. per net per month ...... 56 Figure 9- Mean percent dominance (± S.E.) of all lift nets per month ...... 56 Figure 10-Mean Balanus spp. abundance (± S.E.) per net...... 57 Figure 11-Mean richness (± S.E.) on live oysters compared to disarticulated shells...... 57 Figure 12- Mean percent dominance (± S.E.) on live oysters compared to disarticulated shells...... 58 Figure 13- Mean Balanus spp. abundance (± S.E.) on live oysters compared ...... 58 Figure 14- Mean diversity (± S.E.) of lift nets per month ...... 59 Figure 15-Diversity on live oysters compared to disarticulated shells ...... 59 Figure 16-Mean sediment (± S.E.) load per month from June 2004- June 2005...... 60 Figure 17- Mean silt/clay percentage (± S.E.) per month from June 2004-June 2005...... 60 Figure 18- Mean salinities (± S.E.) of Mosquito Lagoon on lift net sampling dates ...... 61 Figure 19- Mean monthly temperatures (± S.E.) from lift net sampling dates in Mosquito Lagoon...... 61 Figure 20- Historic shell midden sites (=archeological sites) and nearby extant oyster reef sites...... 62 Figure 21- Comparison of size of midden to extant oyster shells ...... 63 Figure 22- Example of an oyster shell covered with barnacles...... 63 Figure 23- Balanus spp. of Mosquito Lagoon. A: B. amphitrite and B: B. eburneus...... 64 Figure 24- Oyster settlement experimental set-up. A: Recirculating flow tank. B: Still water tub ...... 64 Figure 25-Field enclosure for oyster growth and survivorship experiment ...... 65 Figure 26-Oyster settlement (± S.E.) on barnacle densities and species treatments...... 66

vii Figure 27-Mean growth (± S.E.) over 4 weeks...... 67 Figure 28- Cumulative survival of C. virginica over 4 weeks: Trial 1 (October-November 2004) ...... 68 Figure 29- Cumulative survival of C. virginica over 4 weeks; Trial 2 (July- August 2005)...... 69 Figure 30-Sediment and flow rate effect on the settlement of the oyster Crassostrea virginica. Sediment loads were 0 g/ml; 8 g/ml; 16 g/ml...... 70 Figure 31-Sediment and flow rate effect on settlement of the barnacle Balanus amphitrite. Sediment loads were 0 g/ml; 8 g/ml; 16 g/ml...... 70

viii LIST OF TABLES

Table 1- Lift net deployment and collection dates...... 72 Table 2- Total numbers of sessile species recruited to shells in lift nets ...... 73 Table 3- ANOVA comparison of species richness in lift nets...... 75 Table 4- ANOVA comparison of barnacle abundance in lift nets...... 75 Table 5- ANOVA comparison of percent community dominance in lift nets...... 75 Table 6- ANOVA comparison of diversity found in lift nets...... 76 Table 7- ANOVA comparison of total sediment loads collected per month...... 76 Table 8- ANOVA comparison of silt/clay fraction of sediment loads collected per month. 76 Table 9-Shell midden historic sites and closest extant oyster reefs in Mosquito Lagoon ...... 77 Table 10- Separate one-way ANOVAs comparing midden versus extant oyster shell sizes (length and width)...... 78 Table 11- ANOVA to compare settlement of barnacles on historic versus extant shells. Shell source = midden or extant shells...... 78 Table 12-Settlement of oysters on shells with varying barnacle density and species either in flow and still water...... 79 Table 13- ANOVA comparison of oyster growth at the end of four weeks ...... 80 Table 14-Wilcoxon survivorship statistics- Comparison of treatments in Trial 1...... 81 Table 15- Wilcoxon survivorship statistics- Comparison of treatments in Trial 2...... 82 Table 16- Total amount of sediment used in trials ...... 83 Table 17- Sediment and flow experiments: barnacles trials...... 84 Table 18- Sediment and flow experiments: oysters trials...... 85 Table 19-ANOVA comparison settlement of the oyster Crassostrea virginica in varying flow speeds and sediment loads...... 86 Table 20- ANOVA comparison settlement of the barnacle Balanus amphitrite at varying flow speeds and sediment loads...... 86

ix GENERAL INTRODUCTION

Indian River Lagoon System

Human activities threaten productivity, diversity, and survival of coastal resources

leading to a growing need to understand and manage coastal zones (e.g. Jackson et al. 2001).

The Indian River Lagoon system (IRL) was identified as one of the most important and

productive estuarine systems in North America (Fig. 1) (Smithsonian 2001). The Indian

River Lagoon system contains one of the highest species diversities of any estuary in North

America, supporting more than 3000 animal and plant species (Provancha et al. 1992;

Smithsonian 2001). This estuary stretches more than 251 kilometers, from Ponce de Leon

Inlet to Jupiter Inlet, covering over 30% of Florida’s east-central coast (Fig. 1). The Lagoon

system is a series of three distinct, but connected, estuaries: the Indian River, the Banana

River, and Mosquito Lagoon (Walters et al. 2001; Smithsonian 2001).

The northern-most portion of the IRL system, Mosquito Lagoon, lies mostly within

Canaveral National Seashore (CANA). Mosquito Lagoon stretching 30 kilometers and

encompassing 243 square kilometers is a bar-built type estuary bordered by the Atlantic

coastal ridge on the west and the Atlantic beach ridge on the east (Smithsonian 2001). A unique characteristic of the lagoon is its location along the border between temperate and sub-tropical climates leading to a tremendous richness in diversity (Walters et al. 2001).

Over the course of a year, species compositions vary according to the climate. Tropical species dominate during the summer months, while temperate species dominate each winter

(Walters et al. 2001). Consequently, Mosquito Lagoon acts as a refuge for 14 federally listed threatened and endangered species, is nationally recognized for recreational fishing, and is the southernmost U.S. limit along the Atlantic coast for intertidal reefs of the eastern oyster

Crassostrea virginica (Grizzle and Castagna 1995).

1 Populations of Crassostrea virginica within Mosquito Lagoon recently have undergone

significant die-offs (Grizzle et al. 2002). Aerial imagery and field studies have documented

increasing dead zones of disarticulated shells along the seaward edges of the oyster reefs

(Fig. 2) (Grizzle and Castagna 1995; Grizzle et al. 2002). This has resulted in some reefs of

Crassostrea virginica declining over 50% in total area between 1943 and 1995 (Grizzle et al.

2002). Restoration efforts have begun within Mosquito Lagoon and are focusing on

reconstructing the three-dimensional reef habitats (Wall 2004). These efforts hope to re-

establish the ecological role of what are now merely footprints of former reefs. The critical

component here is scientific research. Before the most effective protocol can be established,

important questions about the sources of juvenile and adult oyster mortality must be

answered. Potential causes of C. virginica mortality in the Indian River Lagoon system

include sediment loads, competition, predation, and disease.

Despite being part of a major population center that subjects it to tremendous

stressors from development, recreational use, and introduced species, Mosquito Lagoon and

its natural resources are replenishable. If the nature-driven processes of this system are

sustained, the IRL can continue to function and support the rich variety of plants and

animals it presently contains (Smithsonian 2001). The ecological importance of this area has

been established by the Environmental Protection Agency by listing it as an Estuary of

National Significance and by the State of Florida in classifying it as a Florida Outstanding

Waterway and an Aquatic Preserve (Walters et al. 2001). These designations have supported

both local and national efforts to better protect the biodiversity of this ecosystem and to

further maintain the system in its existing conditions by protecting it against further

anthropogenic effects (Gardner 1991; Smithsonian 2001). The IRL maintains its economic

importance by generating over $ 800 million in revenue annually to the local economy (IRL

2 Comprehensive Management Plan 1996). This includes $300 million income from fisheries

plus $13 million in shellfisheries alone within the IRL (IRL Comprehensive Management

Plan 1996). Within this estuary, C. virginica is one of the most significant economic

contributions and it is harvested recreationally and commercially solely within Mosquito

Lagoon (IRL Comprehensive Management Plan 1996).

In 2001, there were over 90,000 registered boats within the counties that border the

northern IRL system, and this number had increased nearly 10% annually since 1986 (Hart

1994; ANEP 2001). Although many concerns with the increasing number of boaters have

been documented, the impact of rapidly increasing boat activity on important benthic organisms, including the oyster Crassostrea virginica, is a topic now being heavily studied

(Walters et al. 2001; Wall et al. in press). Understanding and quantifying the negative effects of increased water motion and high levels of sedimentation associated with increased boating on larval settlement and oyster survival is critical to determining which mechanism(s) cause oyster reef declines (Grizzle et al. 2002). For example, experiments show that larval settlement of Crassostrea virginica is negatively affected by increased water flow and increased

sedimentation (Shelbourne 1957; Seliger et al. 1982; Nowell and Jumars; 1984; Wall et al. in

press).

Importance of Crassostrea virginica

The economic significance of C. virginica is matched by its ecological importance.

Thus, it is of tremendous importance to include oyster reefs in plans to protect and restore

Mosquito Lagoon. Reefs of Crassostrea virginica are three-dimensional structures created by

years of successive settlement of oyster larvae on adult shells (Dame 1996). Successive

generations increase the structural complexity of the reefs. Its presence in a system has

3 denoted C. virginica worthy of both “keystone species” and “ecosystem engineer” status

(Dame 1996). Crassostrea virginica has been identified as “keystone species” because the removal of these species significantly alters community structure and ecosystem functioning

(Paine 1969). Crassostrea virginica is also an “ecosystem engineer” because it modifies and creates habitats as well as modulates the availability of resources to other species utilizing the three dimensional reef structures (Dame 1996; Coen & Luckenbach 2000).

As the major component of the structural matrix in a system, C. virginica greatly influences its immediate environment (Lenihan 1999). The arrangement of individuals controls the biodiversity of the surrounding community (Bahr & Lanier 1981). Through its complexity, the reefs create varying levels of microhabitats by providing hard surface areas and interstitial heterogeneity that is rare in marine ecosystems typically dominated by soft bottom habitats (Bartol et al. 1999; Micheli & Peterson 1999). Accordingly, its role in habitat creation allows oyster reefs to support more animal life than any other portion of the sea bottom (Nelson et al. 2004). The total number and densities of fish, invertebrate and algal species greatly increase in areas containing oyster reefs (Bahr & Lanier 1981). More than 300 marine invertebrate species may occupy an oyster reef at one time (Wells 1961). In addition to increasing species richness, the three-dimensional structure of the reef provides other services. For example, the established structure stabilizes and buffers shorelines from high wave energy (Smithsonian 2001).

Equally importantly, C. virginica is of ecological significance because of its feeding mechanisms (Coen & Luckenbach 2000; Nelson et al. 2004). Bivalves process materials by consuming particulate and dissolved organic matter and then excreting organic and inorganic nutrients. In this process, they couple the water column to the benthos and change the biogeochemistry of adjacent substrates (Levitan 1995). Crassostrea virginica, therefore, plays a

4 major role in the cycling of nutrients in ecosystems (Nelson et al. 2004). It processes

materials at high rates and speeds up nutrient cycling (Lenihan 1999; Nelson et al. 2004).

Oysters can filter large quantities of water, up to 15,000 times their body volume per one

hour (Loosanoff & Nomejko 1946; Smithsonian 2001). Before the oyster resources were

depleted in the Chesapeake Bay, C. virginica was able to filter a volume of seawater equal to

that of the entire Bay every three days (Newell 1988).

Although beneficial to the community, filter-feeding constantly exposes oysters to

pathogenic microbes as they remove particles and pollutants from surrounding waters

(Nelson et al. 2004). Because oysters are sessile and pump water through their bodies, they are recognized as good ecosystem monitors. Changes in ecosystem health can be noted over time scales varying from hours to years. Because oysters are continually submersed in environmental conditions, they actively contribute to water quality assessments (Smithsonian

2001). In addition, the chemistry of their shell can provide information on global changes in the environment (Surge et al. 2003). Accordingly, oysters have been used as monitors and indicators of stress in marine ecosystems.

5 Biology of Crassostrea virginica

Phylum Mollusca Class Bivalvia Order Pteriode Family Ostreidae

The oyster native to the North American Atlantic coast is Crassostrea virginica, also known as the American or eastern oyster. This species is found within bays and estuarine areas (Gosling 2003). Its range extends from the coasts of Brazil and Argentina northward to the Gulf of Mexico and extending north along the western Atlantic Coast to New

Brunswick, Canada (Burrell 1986; Andrews 1991; Gosling 2003). This oyster is found subtidally throughout its range as well as intertidally in the southernmost parts of its North

American.

General anatomy and physiology

Crassostrea virginica has a highly variable appearance. Its shell consists of two calcareous valves joined by a resilient hinged ligament (Gosling 2003). The shell is usually white and yellowish with purple or brown radial markings (Gosling 2003). It is thick and solid with both valves having concentric sculpting (Gosling 2003). The maximum length for this species is 350 mm, but average length is 89 mm (Gosling 2003). The shape of the shell varies with environment. In an intertidal environment, the shell is thin, irregularly shaped, and elongated, while subtidally the shell is more uniform and thicker (Burrell 1986). The interior of the shell is pearly white with a single posterior purple or black muscle scar (Burrell

1986; Gosling 2003). The lower (left) valve, which is attached to the substrate, is deeply cupped and thicker to accommodate the body (Kennedy et al. 1996). The upper right valve is generally flat and thinner and acts to seal in the body cavity (Kennedy et al. 1996). The

6 oyster settles on its left valve, leading to the asymmetry found in the larger and more deeply

cupped left valve (Kennedy et al. 1996). The large adductor muscle attaches to both shell

valves to control their opening and closing as well as to create a seal when necessary

(Carriker 1996).

Because oysters lack internal skeletons, their calcareous shells serve as an

exoskeleton, providing support for the body, protection for its soft internal organs, and

prevention against the collapse of the mantle cavity (Carriker 1996). The body is made up of

the organs the oyster uses for digestion, respiration, and reproduction (Kennedy et al. 1996).

All of these organs are enveloped in a sheet of tissue called the mantle (Fig. 3) (Kennedy et al. 1996). The mantle joins at the posterior margin of the shell and forms a cap that covers the mouth and labial palps (Kennedy et al. 1996). The labial palps lie at the hinge edge, are flat, and have three papillae folds or lips. The gills are located below these palps and consist of four demibranches, folds of crescent-shaped tissues that occupy most of the mantle cavity

(Fig. 3) (Eble & Scor 1996).

Together with mantle, the gills are the chief organs of respiration (Eble & Scor

1996). Covering the gills are tiny hairs, which create a current for incoming water. This current moves food particles to the labial palps and is located above the gills for sorting and oxygen absorption (Gosling 2003). The mantle is lined with thin blood vessels, which extract oxygen to pass to the three-chambered heart located underneath the adductor muscles

(Gosling 2003). The heart pumps colorless blood to all of the body. The kidney, which is located by the adductor muscles, then purifies the blood of all wastes (Carriker 1996).

Oysters ingest seston (planktonic material) that includes bacteria, protozoa, and a wide variety of phytoplankton, organic detritus, and inorganic material (Quale 1969). The ingested particles are filtered by the gills, then entrapped and bound in mucus. These strings

7 of mucus carry the particulate matter to the ciliated labial palps where it is sorted (Kennedy

& Breisch 1980). Oysters differentially select particles to consume or reject since stomach contents do not reflect all the phytoplankton to which they have been exposed (Korringa

1952). Oysters can filter particles smaller than 2 µm but retention efficiency increases with size (Fritz et al. 1984). Accepted particles are passed from the mouth down the esophagus into the stomach, while rejected particles are secreted as pseudofeces (Kennedy & Breisch

1980; Kennedy et al. 1996). Oysters use an organ unique to bivalves and gastropods called a crystalline style to assist in the digestion process (Galtsoff 1964; Quale 1969a). After digestion, the wastes pass down the intestines into the exhalent chamber where they are stored until release (Menzel 1991). The mechanical process of filter feeding in adult oysters has been well documented by Galtsoff (1964).

The reproductive system in bivalves is a simple pair of gonads made of branching tubules and gametes are budded off the epithelial lining (Gosling 2003). Crassostrea virginica is a protandric species. Generally, when they first mature, all eastern oysters are males

(Galtsoff 1964; Burrell 1986; Mackie 1984). As the oysters grow, however, a portion of each size class becomes females. This results in an excess of females being present in larger age classes (Galtsoff 1964). Adult oysters are usually dioecious, but sex changes and reversals are frequent. Since sex reversals are common, females may become males again and vice versa (Galtsoff 1964; Burrell 1986). There is no way to differentiate between males and females by looking at their shells.

Environmental requirements

Due to their sessile nature, oysters are unable to move when faced with unfavorable environmental conditions. Thus, for the eastern oyster to be a successful estuarine animal, it

8 must tolerate a wide a wide range of physical conditions including varying temperatures,

salinities, water currents, oxygen levels, chemical substance concentrations, and turbidities

(Burrell 1986; Andrews 1979).

Temperature

Temperature not only sets the limits of spatial distribution, but also affects many

aspects of oyster biology, including feeding, reproduction, growth, respiration,

osmoregualtion, and parasite disease loads (Gosling 2003). Eastern oysters are able to

survive in water temperatures between 10° and 43°C (Burrell 1986). This range includes

exposure to low water temperatures and high air temperatures when exposed at low tide

(Burrell 1986). Oysters need a temperature of at least 19.5°C for egg development, above

20°C for proper larval development, and between 10° and 30°C for adult growth (Galtsoff

1964; Burrell 1986). Temperature also impacts feeding. Water movement through the

oyster increases as temperatures rise; consequently, at higher temperatures more available

food is filtered through the gills (Quale 1969). Temperature also has an indirect effect on

oysters by influencing the production of available food (Quale 1969).

Salinity

Eastern oysters are able to live in a wide range of salinities, specifically between 0 and

42.5 ppt (Ingle & Dawson 1950). The open ocean is 32-38 ppt, with an average of 35 ppt

(Gosling 2003). The optimal salinity for C. virginica is 25-28 ppt (Gosling 2003). As with temperature, certain life stages have different salinity requirements and optimal ranges. Egg cleavage occurs between 7.5 and 35 ppt, with optimal development between 10 and 22 ppt

(Castagna & Chanely 1973). Larval development occurs between 5 and 39 ppt, but optimally at 25-29 ppt (Castagna & Chanely 1973). Metamorphosis has been demonstrated to be

9 limited to areas with 5.6-35 ppt seawater (Castagna & Chanely 1973). Spat growth is best at

15-26 ppt (Chanely 1957). Favorable adult growth has been documented from 14-30 ppt, stunted at 7.5 ppt and nonexistent at 5 ppt (Castagna & Chanely 1973).

Oysters respond to salinity changes by controlling their degree of shell opening

(Galtsoff 1964). Thus, salinity plays a big role in the volume of water filtered and the feeding of oysters. When exposed to sudden reduction in salinity from 27 ppt to 5 ppt for

24 hours, there was a 24% decrease in filtration rate when returned to 27 ppt (Loosanoff

1953). Furthermore, while at 5 ppt there was a 100% decrease in filtration rate (Loosanoff

1953). Normal pumping resumed with no long-term effects, immediately (Loosanoff 1953).

Water circulation

Due to the sessile nature of the oyster, water motion plays a principal role in

providing the conditions for feeding and purification, as well as for successful reproduction

and dispersal of oyster larvae (Nelson et al. 2004). Water currents must be strong enough to

provide sufficient food (Burrell 1986). To do this the water must be replenished at least 72

times in a 24-hour period (Galtsoff 1964). Additionally, water circulation is needed to wash

away the biodeposits of animals residing within a reef (Galtsoff 1964; Haven & Morales-

Alamo 1966). A tidal current between 5 and 66 cm/sec has been shown sufficient for this

(Wells 1961).

Too much water motion can also be detrimental. Currents too great can interfere

with feeding and cause structural damage (Galtsoff 1964; Burrell 1986). Excessive water

motion can tumble oysters about, knocking off the fragile edges of shells and causing

structural damage within the reef (Galtsoff 1964; Burrell 1986). Currents of 15 cm/sec

caused unattached shells to tumble along the bottom of Long Island Sound (MacKenzie

10 1979). High wave action also increases water turbidity by stirring up bottom sediments and

causing silt accumulation over the oysters (Korringa 1952).

Chemical effects of water

Kennedy and Bresich (1981) reviewed literature dealing with water chemistry and

organic/inorganic compound effects on the eastern oyster. They reviewed the effects of pH, chlorine, heavy metals, petroleum, hydrocarbons, and detergents on the Crassostrea virginica

and found the various life stages of oysters have different susceptibilities to pollution

(Kennedy & Bresich 1981). Pollution can lead to the loss of oyster larval food supply, poor

larval growth, loss of larval vigor, increased susceptibility to disease, pests or predation,

contamination of settling surfaces, decreased fecundity, and reduced spawning. This is

important because estuaries act as sinks that gather these contaminants and pollutants before

they reach the open ocean (Walters et al. 2001). These various types of pollutants also pose

threats to humans as they can become concentrated into oyster body tissues posing harm

during consumption by higher predators (Gosling 2003).

Turbidity

Oyster settlement and recruitment on oyster beds is greater where shells are

abundant and silt deposits and fouling organisms on shells are scarce (MacKenzie 1983).

Suspended bottom sediments can cause oysters to either stop feeding or expend

considerable energy in separating mud and sand from edible particles (Quale 1969). In

addition, MacKenzie (1977) and Gunter (1979) found that oyster beds covered with a layer

of sediment several centimeters thick and covering less than 2.5 cm on the surface of the

shells reduced oyster settlement. This field study compared reefs in Mississippi where

predation in concert with high sediment and silt levels may have contributed to the decline

11 of recruits. It was hypothesized that the settling larvae were killed by abrasion (Gunter 1979).

While more tolerant of turbidity, adult oysters do show a decrease in pumping rates at silt concentrations above 1.0 g/L (Davis & Hindu 1969). Fertilized eggs experienced 20% mortality at silt concentrations of 0.25g/L (Davis & Hindu 1969). At 0.75 g/L silt concentrations, the growth of larvae was significantly reduced (Loosanoff & Tommers 1948;

Loosanoff 1962; Burrell 1986). Within the Indian River Lagoon, oysters were not efficiently able to settle at sediment concentrations above 8 g/L (Wall 2004).

Reproductive physiology Oysters are stimulated to ripen their sexual organs by cues in the surrounding water.

The cue of foremost importance is water temperature. It stimulates gametogenesis and spawning in the oyster (Burrell 1986). The critical temperature required for spawning varies by location (Burrell 1986). Spawning may be year-round, where warm conditions and sufficient food is continuously available (Burrell 1986; Kennedy et al. 1996). In the Indian

River Lagoon system, there have been some reports of year-round spawning (Grizzle &

Castagna 1995; Smithsonian 2001). Wall (2004), on the other hand, found spawning to only occur from late spring through the fall in some years (Wall 2004). Besides temperature, another physical factor important for spawning is salinity.

Under natural conditions, gametes from the males are released first, and then egg release is immediately triggered in females (Galtsoff 1964; Bahr & Lanier 1981). Oysters release gametes by gently clapping their shells together. Galtsoff (1964) has thoroughly described gametogenesis and histology of developing ova and sperm.

External fertilization takes place in open water column with the first cell division occurring within 6 hours (Burrell 1986). Fertilized eggs are pear-shaped and multilayered with jellylike membranes (Galtsoff 1964; Burrell 1986). They range from 55-75 mm in

12 lenght and 35-55 mm wide (Galtsoff 1964). A detailed description of the anatomical

development of C. virginica larvae from first cleavage to settling is provided by Galtsoff

(1964).

Larvae and settlement

If conditions are favorable, larvae reach the trochophore stage within 12 hours of fertilization (Fig. 4) (Burell 1986). This is followed by 90-95% of the fertilized eggs developing to the shelled veliger stage within 48 hours, in hatchery conditions (Burrell 1986).

Because the larvae are not capable of metamorphosing immediately after fertilization, the resulting larvae are planktotropic (Burrell 1986). During this planktonic life stage, the oyster larvae move with the currents, feed on the plankton, and grow over a period of two to three weeks (Bahr & Lanier 1981). At this time, mortality rates can exceed 70% (Rumrill 1990;

Dame 1996).

Oyster larvae are typically concentrated near the water’s surface during rising tides and near the bottom during falling tides (Sellers & Stanley 1984). This decreases their chances of being more widely distributed and carried offshore (Carriker 1951). Therefore, a primary determinant of the distribution of a pelagic planktonic species is the prevalent water currents during the critical free-swimming stages of the larvae. These hydrodynamic regimes ultimately control the number of larvae that are brought into the vicinity of settlement substrate (Connell 1985; Gaines et al. 1985).

The growth and length of the larval period is dependent on the water temperature and food supply, with greater growth associated with higher concentrations of food and higher water temperatures (Burrell 1986). Lower temperatures and food supplies prolong development and pelagic existence (Underwood & Fairweather 1989). Any increase in time

13 in the larval stage undoubtedly leads to decreased survival due to increased exposure to

predators (Underwood & Fairweather 1989). The final larval growth stage in oysters is

called the pediveliger, or eyed larva (Fig. 4). Pediveligers reach 300 µm in length (Burrell

1986). A well-developed foot and two eyespots, which serve as a defense mechanism, enable

the larvae to crawl on the bottom in search of substrates suitable for attachment (Burrell

1986).

It was long thought that dispersal and settlement of marine invertebrate larvae was a

random process caused by the tides and currents with only a small portion of the larval stock

surviving and reaching appropriate substratum by chance (e.g. Underwood & Fairweather

1989). It is now known that larvae can delay settlement for extended periods up to three

weeks and allow for a longer searching phase with a greater probability of finding the

optimum site (Lewis 1978).

Once competent to settle, oyster larvae swim to the bottom and crawl about using

their velum, gradually restricting their search area until a final attachment site is found

(Cranfield 1973). While searching for suitable substrate, larvae may respond to physical

factors received through surface mechanoreceptors (Hadfield 1978). They distinguish

between vertical and horizontal surfaces by statocysts and distinguish lighted surfaces by the

shading received by larval eyes (Hadfield 1978). This allows them to distinguish factors

critical to settling that include sufficient light and surface irregularities on the substrate

(Galtsoff 1964; Ritchie & Menzel 1969). Larvae also respond to chemosensory cues. These

cues can be found on the substrate itself and or in water solution, which can be sensed

without the larvae actually encountering the substratum (Turner et al. 1994). Larvae of C. virginica are capable of discriminating between substrates, as evidenced by their behavior before attachment. The presence of oyster shells is typically enough to induce settlement

14 (Crisp 1967). Bacterial films on the surface of oysters shells, pheromones released by live oysters, and various metabolites of oysters all act as attractants for settling larvae (Hadfield

1978; Coon et al. 1985,1990 a,b; Fitt & Coon 1992). Examples of chemical cue inducing oyster settlement include the neurotransmitters L 3,4 dihydroxyphenlyalanine (1 DOPA), epinephrine (EPI), and ammonia (NH3) ( Hadfield 1978; Coon et al. 1985,1990a,b; Fitt &

Coon 1992).

Post-Settlement

Once a suitable substrate is found, the larva responds to metamorphic stimuli by

becoming benthic and losing their velum (Crisp 1976; Hadfield 1978). They do this by

turning onto their left side, ejecting cement produced by the byssus gland, and affixing themselves to a hard surface (Burrell 1986). Most often, larvae choose conspecific oyster shells for settlement (Burrell 1986; Kennedy et al. 1996; Gosling 2003). At this time, they undergo metamorphosis and are called spat (Kennedy et al. 1996). Pediveligers rapidly lose larval features after attachment (Galtsoff 1964). This metamorphosis from free-swimming

larvae to spat is accompanied by some marked morphological changes, including the

disappearance of the velum, foot and the anterior adductor muscle, and the development of

an enlarged set of gills (Loosanoff 1965). Eyespots are lost and the foot and velum are

incorporated into parts of the alimentary systems (Galtsoff 1964). After settling, the juvenile

oyster is a sessile animal and will remain immobile at this attachment site for life (Burrell

1986). Unable to move around, the oysters cannot move away from predators, competitors,

or relocate to areas with more phytoplankton (Zimmer-Faust & Tamburri 1994).

15 Growth

Oysters feed on plankton in the water column. When the water conditions are

optimal and the nutrients are sufficient, oysters feed continuously (Sellers & Stanley 1984).

Growth in oysters varies widely with tidal height, growing area, and environmental conditions, but is proven greatest right after settling and decreases with age (Burrell 1986).

Under the right conditions, sexual maturity can be reached in as little as four weeks (Cake

1983). Oyster growth is typically measured by an increase in shell size or body size. Growth

is continuous throughout the year as far north as South Carolina, although it slows in winter

months (Ingle & Dawson1950). Florida oysters in Apalachicola Bay have been documented

to grow 100 mm in 31 weeks (Burrell 1986).

The growth of oysters is slower on intertidal reefs than in subtidal areas due to

intraspecific crowding experienced in the intertidal beds and reduced feeding times (Burrell

1982). Interspecific fouling organisms associated with oysters also have significantly negative effects on growth, even causing growth to cease altogether (Osman et al. 1989,

Zajac et al. 1989).

Mortality factors

Parasites and Diseases

In recent years, the importance of oyster mortality due to diseases and parasites has

been a major research focus. A complete description of reported disease and mortalities of

Crassostrea virginica can be found in Ford and Tripp (1996) and Gosling (2003). Only the

most common parasites and diseases are presented here. Protozoans are responsible for

both Dermo and MSX diseases. Infections of Perkinsus marinus cause Dermo. It is present

both in subtidal and intertidal oysters along the Atlantic coast south to Venezuela and

16 throughout the Gulf of Mexico (Burrell 1986). Dermo causes emaciation, the loss of body

condition, and is easily spread from oyster to oyster. This leads to extremely high moralities,

100%, in some locations in recent years (Ford & Tripp 1996). Other diseases, such as MSX

(Minchinia nelsoni) and SSO (M. costalis), have not been reported south of North Carolina on the Atlantic coast (Gosling 2003). These diseases also cause emaciation, reduced health, and fecundity, and can result in 100% mortality. Parasites, such as Bucephalus cuclis, are

trematodes that infect the gonads, gills, and the digestive gland (Ford & Tripp 1996). The

result is castration with an associated mortality of 30%. Certain infections are exclusive to

larva. In the hatchery the bacteria, Vibrio spp., only affects larvae and causes 100% mortality

(Ford & Tripp 1996). Additionally, a new species of alpha proteobacteria (CVSP) causes

juvenile oyster disease (JOD) (Gosling 2003). It occurs only in oysters less than 25 mm,

resulting in over 90% mortality (Gosling 2003).

Predators

Predators play various roles during the life-cycle of C. virginica (e.g. Osman et al.

1990). During the planktonic egg and larval phase, mortality is high due to predation from

larger organisms feeding on the plankton (Gosling 2003). These organisms include

ctenophores, adult bivalves, anemones, starfish, fish, and crustaceans (Gosling 2003). Mud

crabs, juvenile blue crabs, and flatworms typically eat the newly settled spat (Gosling 2003).

Blue crabs, whelks, boring sponges, oyster drills, rays, and several species of fish typically eat

adult C. virginica (Walters et al. 2001). Together, these predators limit the growth and

survivorship of C. virginca. Intraspecific predation between size classes is also frequent as post-settlement conspecifics consume larvae (Gosling 2003). In doing so, adults remove

17 potential competition for food, space, and other resources (Underwood & Keough 2001;

Dame 1996).

Competition

Competition for space and food plays an important role for C. virginica. The ultimate result is a reduction in the number of oysters, a change in recruitment patterns influencing community structure, and a reduction in physical condition (Lenihan & Micheli 2001).

Organisms that compete with oysters for space and food are mainly encrusting organisms and include the following: macroalgae, sponges, cnidarians, bryozoans, barnacles, ascidians, anemones, polycheates, mollusks, and arthropods (Wells 1961; Gosling 2003). The main effects of competitors are: 1) prevention of settlement by coverage of available space, 2) allelochemicals that deter settlement of new recruits, and 3) overgrowth or poisoning (White

& Wilson 1996). Of the four main effects of competition, restricted space for settlement and overgrowth is the most common (Osman et al. 1989). Typically, the distribution of intertidal oysters does not appear to be entirely suppressed by the presence of the interspecific organisms (Burrell 1986). Nevertheless, even when the oysters are not killed, competitors cause reduced growth and survival in oysters (Zajac et al. 1989). One such example of a competitor is the slipper shell gastropod, Crepidula, which is the most important competitor to Crassostrea virginica in Europe (Gosling 2003). It is so successful because it settles around the same time as the oyster and it grows more rapidly than the oysters, causing mortality rates of up to 60% (Gosling 2003).

18 Crassostrea virginica in Mosquito Lagoon

Crassostrea virginica is the primary oyster species that inhabits the Mosquito Lagoon

(Smithsonian 2001). Although the ecological importance of oysters has been known for a long time, they have historically been treated as a species to exploit rather than conserve

(Micheli & Peterson 1999). Recently, concern for this species has spread throughout the entire east coast of the United States (Micheli & Peterson 1999). Increasing anthropogenic disturbances along the coast have led to substantial losses in eastern oyster populations

(Coen et al. 1999). Threats include overfishing, disease, and habitat degradation (Coen et al.

1999). The economic and ecological importance of this species therefore calls for efficient approaches to the conservation and management of wild populations. It is vital to protect and restore these refuges of biodiversity. Because Mosquito Lagoon is the southernmost limit on the Atlantic coast for undisturbed intertidal reefs of the eastern oyster, it is thus an area of extreme importance (Grizzle & Castagna 1995).

19 CHAPTER ONE:

BIODIVERSITY OF SESSILE SPECIES ON OYSTER REEFS

Introduction

Three-dimensional reef structures of Crassostrea virginica are created by years of

successive settlement of oyster larvae onto adult shells (Dame 1996). Successive generations

increase the structural complexity of the reefs denoting C. virginica worthy of both “keystone species” and “ecosystem engineer” status (Dame 1996). Through its structural complexity, the reefs create varying levels of microhabitats by providing hard surface areas and interstitial heterogeneity that is rare in marine systems dominated by soft bottom habitats (Bartol et al.

1999; Micheli & Peterson 1999). Accordingly, oyster reefs support more animal life than any other portion of the adjacent sea bottom; more than 300 marine invertebrate species and numerous other taxa, including fishes, macroalgae, etc., have been found associated with oyster reefs (e.g. Wells 1961; Bahr & Lanier 1981; Nelson et al. 2004). Reef structures also stabilizes and buffers shoreline from high wave energy, influences water currents, changes hydrodynamic regimes, and reduces organismal exposure to hydrographic conditions (e.g.

Bahr & Lanier 1981; Micheli & Lenihan 1999; Nelson 2004).

Populations of Crassostrea virginica within Mosquito Lagoon recently have undergone significant die-offs (Grizzle et al. 2002). Some reefs of Crassostrea virginica have declined 50% in total area from 1943 to 1995 (Grizzle et al. 2002). Aerial imagery and field studies have documented that reefs along major boating channels have dead zones of disarticulated shells,

“dead margins”, along their seaward edges due to boat wakes (Fig. 2) (Grizzle & Castagna

1995; Grizzle et al. 2002). Restoration efforts have begun within Mosquito Lagoon and are focusing on reconstructing the three-dimensional reef habitats (L. Walters pers comm.).

20 These efforts are intended to re-establish the ecological role of what are now merely

footprints of former reefs. The critical component here is scientific research. Before the

most effective protocol can be established, a better understanding of the ecology of these

reefs must be established.

This study investigates the influence of dead margins on the recruitment of sessile

organisms on oyster reefs in Mosquito Lagoon. Few studies have looked specifically at the

diversity of species on oyster reefs (e.g. Wells 1961, Crabtree and Dean 1982, Wenner et al.

1996, Coen et al. 1999). This is the first study in Florida to specifically look at organisms

that permanently attach to oyster shells. Lift net methods were adapted from studies

conducted on fishes and decapods by Tolley et al. (2005) on the west coast of Florida.

Additionally, I compared sessile invertebrate recruitment in Mosquito Lagoon on extant

reefs to preserved oyster middens to determine if sessile inhabitants have changed over the

course of the past 15,000 years.

Methods: Lift Nets (Present Day Species)

Study Site

Research was conducted in Mosquito Lagoon, within the boundaries of Canaveral

National Seashore (Fig. 1). Mosquito Lagoon is the northeast estuary in the Indain River

Lagoon system in a series of three distinct, but connected, estuaries which extend 251

kilometers (156 miles) from Ponce de Leon Inlet to Jupiter Inlet on the east coast of central

Florida. The average depth of the Lagoon is less than 1 meter in most areas and the current

is primarily wind-driven (Walters et al. 2001). Annual salinity ranges between 18 and 45 ppt,

depending on rainfall (Grizzle 1990; Walters et al. 2001; M. Boudreaux unpublished data).

21 Most of the lagoon is a complex system of shallow and open water areas with nearly 100 mangrove (Rhizophora mangle and Avicennia germinans) dominated islands (Walters et al. 2001).

Intertidal oyster reefs are found throughout this region.

Lift Net Field Sampling and Analysis

Ms. Jennifer Stiner, a graduat student at the University of Central Florida, and I collaborated extensively on this project. She collected data on mobile species while my research focused on the sessile species

(flora and fauna). I present only my results here.

Six oyster reefs were selected for this study, three in pristine condition, and three that have dead margins (Figure 5). Five replicate lift nets were placed on the backreef area of each oyster reef. The protected backreef areas were chosen to minimize the loss of nets due to wave action. Lift net methods were taken from Crabtree and Dean (1982), Coen et al.

(1996), and later modified by Tolley et al. (2004) for use in Florida systems. I further modified the protocol to include the enumeration of sessile species occupying oyster reefs.

I created 0.5 m2 lift net frames from 3.75 cm diameter PVC; nets were 0.5 m deep

(Fig. 6). The sides of the net were made from 3.2 cm diameter opening mesh and the bottom was made from a 1 m square of 0.2 cm diameter opening mesh. I machine sewed the two mesh sizes together using cloth extra strength thread. I then attached the sewn mesh to the PVC frame with cable ties (300 mm in length; weight limit: 11 kg) to construct the nets.

I deployed all nets intertidally, just above mean low water, on living oyster reefs (Fig.

6). Volume normalized oyster shells (1.5 L) were placed in the lift nets. Approximately half of the shells (0.75 L) were single, disarticulated shells and half were live clusters. All were scraped clean of all epiflora and epifauna. New shells were placed into the nets each month.

Additionally at time of net retrieval all nets were brushed clean to remove organism that had

22 settled upon the nets or PVC frames. Organisms recruiting to shells in lift nets were collected by swiftly picking up nets on two sides and retrieving contents (Fig. 7). I collected the lift nets monthly for 12 months (June 2004-July 2005) (Table 1). An extra month was added as the 2004 hurricane season prevented September 2004 data collection. Thus, the nets were deployed for a total of 13 months. During each collection, all net contents were brought back to the lab where I identified all sessile organisms within 24 hours and returned them alive to the lagoon.

Environmental Variables

Permanent temperature monitors (Onset Stowaway Tidbit Temperature Loggers) were deployed at each site attached to cinderblocks placed at water depth equal to the net depth. Hourly temperature was recorded for the entire 13 months. I measured salinity upon net retrieval using a portable refractometer (VeeGee A366ATC). Three sediment traps were deployed at each site at the same depth at the lift nets to determine sediment load accumulations during the 4-weeks intervals between sampling. Each replicate, cylindrical

PVC pipe sediment trap (10 cm diameter X 25 cm deep) was submerged flush with substrate

(Lenihan 1999). I capped traps underwater at the time of retrieval. I retrieved the sediment traps concurrently with the retrieval of lift nets and new traps were immediately deployed to replace them. I determined the total sediment mass by drying samples at 60° C for 48 hours in a drying oven (Econotherm Model Number 51221126) and weighing contents on a top loading balance (O’Haus Scout 2-Model Number SC6010). Relative grain size was determined by grinding the dried sediment and sorting samples with a sieve (0.062 mm) to separate the silt/clay from the sand/grain fractions.

23 Statistical Analysis

Community metrics of sessile species recruitment were examined. Response variables of species richness (total number of species), dominance (percent occurrence of most abundant species), species diversity (Shannon-Weiner), and the abundance of the most important taxa, were analyzed using a three-way, nested analysis of variance (ANOVA).

Prior to running all ANOVAs, homogeneity of variance and normality were tested. Fixed factors in the nested ANOVAs were: 1) reef type (dead margins or unimpacted), 2) month, and 3) shell type (settlement on dead versus live oyster shells). Shell type was nested within each site. Site was nested within reef type.

A two-way ANOVA was conducted to test whether sediment loads on oyster reefs varied as a function of the following fixed factors: reef type (dead margins or unimpacted) and month. Means per reef per reef type were calculated.

Results

Biodiversity and Composition

Twenty-four sessile species of sessile invertebrates recruited to oyster shells were collected in the lift nets during my study (Table 2). Balanus species (Arthropoda) dominated all samples numerically. Mollusca represented the most abundant phyla, with nine species found. Others were classified within Annelida, Cnidara, Porifera, Ectoprocta, and Chordata

(Table 2). Measures of oyster community metrics exhibited clear trends in Mosquito Lagoon.

There was no significant effect of the replicate sites on any of the response variables (species richness, dominance, species diversity, or the abundance of most important taxa).

Alternatively, the community metrics of species richness, dominance, and the abundance of

24 most abundant taxa (Balanus species), differed temporally, due to the month of sampling

(p<0.001) (Figs. 8,9,10; Tables 3,4,5). Richness was significantly higher during June, July,

August, and October (p=0.007); (Fig. 8). Additionally, February had the lowest richness

(p=0.001) (Fig. 8). Community percent dominance (% occurrence of most dominant

species) was significantly greater in August and February (p=0.012) (Fig. 9). These samples

were dominated by Balanus spp. Balanus species abundance was significantly higher in June,

July, and August (p=0.003) (Fig. 10). Furthermore, species richness, dominance and the

abundance of Balanus spp. were all higher on living oysters than on single disarticulated

oyster shells (p<0.001) (Figs. 11, 12, 13). Diversity however did not show any significant differences, species diversity did not differ among the sampling periods (months) or the shell type (live clusters versus disarticulated shells) (p=0.095 and p=0.159 respectively) (Figs. 14,

15; Table 6).

Sediment Loads

A two-way ANOVA was conducted to test whether sediment loads on oyster reefs

varied as a function of reef type (impacted or unimpacted) or month. No significant

difference was observed between the total amount accumulated on unimpacted oysters reefs

versus those with dead margins (p=0.872) (Table 7). neither the replicate site nor the

sampling period (month) had an effect on the sediment accumulations (p=0.320; p=0.198);

(Fig. 16; Table 7).

Percentage of the silt/clay fraction of the total sediment load was also calculated

(Table 8). No significant difference was observed between the total amount accumulated on unimpacted oysters reefs versus those with dead margins (p=0.504) (Fig. 17; Table 8 ).

25 However, the month of sampling was significant in the percentage of the total sediment that was silt/clay (p<.001) (Fig. 17; Table 8).

Environmental Factors

Salinity ranged from 25 to 35 ppt in Mosquito lagoon during this study (Fig. 18). The lagoon normally averages 34 ppt but lower salinities were attained due to significant rainfall the 2004 hurricane season. Salinity did not differ among sites or sampling periods.

Temperatures ranged from 16° C in February 2005 to 31° C during the course of study (Fig.

19).

Methods: Middens (Historical Analysis of Organisms Attached to Oysters)

Numerous introduced species of macro invertebrates have been reported from

Mosquito Lagoon. Thus it was important to examine the effect invasive species may have had on the utilization of the eastern oyster as settlement substrate. To determine if the diversity and absolute abundance of sessile organisms attaching to Crassostrea virginica has changed, I examined 244 historical disarticulated shells from the Mosquito Lagoon and compared them to 300 shells from extant reefs in Mosquito Lagoon. The historical shells were taken from three archeological regions in northern Mosquito Lagoon within the boundaries of Canaveral National Seashore: 1) Oyster Bay, 2) Turtle Mound, 3) and

Seminole Rest (Fig. 20; Table 9). The dates for the midden shells have been determined by radiocarbon dating (Table 9). Extant shells were collected from the closest living reefs to the archeological sites (Table 9).

26 All historic shells were recovered by the National Park Service archeological teams,

desalinated and preserved in airtight containers at the Southeastern Archeological Center, in

Tallahassee Florida. I visually inspected 244 disarticulated shells of C. virginica to quantify the

numbers of sessile species, present on the preserved shells. Specifically, the following

parameters were measured on each shell: maximum length and width of shell, number and

size of juvenile oyster spat, number, size and species of identifiable intact Balanus shells,

number and size of barnacle scars, and any other species present on each shell. The distance

from each species to oyster spat present was also recorded. A two-way ANOVA was used to

compare the numbers of Balanus spp. found on historic shells to extant shells from Mosquito

Lagoon. The factors in the ANOVA were site (Oyster Bay, Turtle Mound, or Seminole Rest)

and age of shells (historic or extant). Separate one-way ANOVAs were also conducted to

compare shell length and width for the historic to extant oysters.

Results

Similar sessile species were found on midden shells and the living reefs within the

study area. Species found on the midden shells included the barnacle Balanus eburneus, the polycheate worm Hydroides sp., the boring sponge Cliona sp., and the bryozoan Conopeum sp.

All of these species were present on extant shells. Additionally, the extant reef had settlement of the slipper snail Crepidula sp., the jingle shell Anomia simplex, and the invasive purple striped barnacle Balanus amphitrite. The mean size of extant oysters shells, was not significantly different from the midden shells, at any of the three sites examined (ANOVAs: length p=0.443; width p=0.789) (Fig. 21; Table 10). However, there was a significant difference between the numbers of barnacles (intact and scars) on historic versus extant shells (Table 11). The mean number of Balanus eburneus per shell from the middens was

27 0.3548 + S.E. and the average number of barnacles (B. ebernues and B. amphtirite) on living

reefs was 1.5078 + S.E. (p< 0.001). Turtle Mound had a higher abundance of Balanus spp.

than any other site both extant and in midden shells (p=0.008) . Few midden specimens

contained any other organisms (e.g. oyster spat, Hydroides sp., Cliona sp., and Conopeum sp.) so analysis were not conducted on these species.

Discussion

The assemblage of sessile species collected in association with the oyster reefs during the lift net study was similar to those previously reported with oyster reefs (Wells 1961 subtidal in North Carolina). These data are also supported earlier research conducted in the

Indian River Lagoon system that looked at the sessile species diversity on hard substrata, although not specifically associated with Crassostrea virginica (Mook 1976, 1980, 1981, 1983).

Many established exotic species were identified within the Lagoon including B. amphitrite, which was established approximately 100 years ago (Table 2) (J. Carlton pers comm.). A new exotic mussel species not previously recorded to be in the Indian River

Lagoon was also discovered during the course of this research and was identified as Mytella charruana (Boudreaux & Walters submitted).

Dead margins occurring on reefs do not seem to have a significant effect on the backreef usage of oysters as substrate by sessile species. The backreef areas were similar on pristine reefs and reefs with dead margins. This suggested the backreef area on oyster reefs with dead margins still function similar to a pristine oyster reef. Additionally, sessile organisms prefer to settle on living oysters rather than on dead disarticulated shells. This is most likely due to the accumulation of biofilm that was present on living shells as opposed to dead which lacked the biofilm initially. These shells would have lacked chemical cues from

28 adult oyster that induce larval settlement (Tamburri et al 1996). Suffiecent biofilm would be present with a few days of placing shells in the field (Tamburri et al. 1996). Furthermore, the three-dimensional structure of the two different settlement substrates was very different.

Disarticulated shells were single and loose, while the live shells were attached together in clusters. These clusters may have provided more protection and refuge from predators for sessile inhabitants.

In previous research in Mosquito Lagoon, Wall et al. (in press) found an increase in sediment accumulation on the seaward edges of reefs with dead margins. Wall et al. (in press) suggested this was due to resuspension by boat wakes. Moreover, increased sediment has been shown to decrease the survival of newly recruited Crassostrea virginica in these locations (Wall 2004). Thus, any difference in sediment loads between sites would have been predicted to have an effect on sessile species assemblages between the two types of reefs

(impacted vs. unimpacted). This study on the backreef regions of these same reefs did not show any differences in sediment loads between reef types. In contrast, Wall (2004) found that reefs with dead margins had an increase in sediment load on the outer edges of impacted reefs as compared to unimpacted reefs. The dead margins of the reefs are hypothesized to protect these backreef areas preventing sediment accumulation.

Presence of sessile species on historic shells from middens and extant shells in

Mosquito Lagoon was similar and included the barnacle Balanus eburneus, polycheate worm

Hydroides sp., boring sponge Cliona sp., and Conopeum sp. Furthermore, the extant reef counterparts had settlement of the slippersnail Crepidula sp., the jingle shell Anomia simplex and the invasive barnacle Balanus amphitrite. The lack of Crepidula sp. and Anomia simplex in the midden samples is not surprising as these species do not leave defined permanent marks and may be hard to discern in this type of study. More notable was the appearance of

29 Balanus amphitrite on extant shells. Balanus amphitrite originated from the Southwestern Pacific and Indian Oceans (Zullo 1963). Its range now extends from Cape Cod to the Caribbean.

Its spread is attributed to ship fouling (Zullo 1963). It is a recent invader to the IRL within the past 100 years (J. Carlton pers comm.). Mean total barnacle abundance was 5X higher per shell on extant reefs than on midden shells. As the total number of these sessile species has increased, on interference competition has undoubtedly also increased in the area. These data are corroborated with accounts of Balanus spp. monopolizing the settlement space of

Crassostrea virginica on earlier oyster recruitment trials in Mosquito Lagoon (Wall et al. in press).

Diversity is extremely high in the IRL due to its location within a zoogeographic transition zone (Walters et al. 2001). Biodiversity measurements tell us about the interactions between ecosystem composition, structure, and functioning. My study documented histories and current biodiversity associated with oyster reefs. These data provide a baseline from which to evaluate efforts to practice sustainable ecosystem management of Mosquito

Lagoon.

30 CHAPTER TWO:

EFFECT OF BARNACLES ON OYSTER SETTLEMENT, GROWTH AND

SURVIORSHIP

Introduction

Sessile marine organisms share two common resource pools: the substratum on which they attach and the aqueous environment, the source of required physical and organic nutrients (Lohse 2002). Lack of these resources or increased consumers compared to the

availability of resources results in density-dependent competitive interactions (Underwood &

Keough 2001). Competitors may physically interfere with settling species or deprive others by utilizing food resources (e.g. Buss 1979; Buss & Jackson 1981; Okamura 1986; Frechette

& Aitka 1992; Lohse 2002). Competition influences the sizes, fecundities, and densities of all species involved (e.g. Underwood & Keough 2001). Those species that are able to monopolize the resources will successfully be able to limit other species (Bell 2003).

Studies have shown the mortality of newly settled sessile species, including oysters and barnacles, to be due to overgrowth, dislodgement from the substrate, and local food depletion by co-occurring species (e.g. Buss and Jackson 1981; Keough & Downs 1982;

Davis 1987; Peterson & Black 1987; Osman et al. 1989; Bell 2003; Gosling 2003). For example, studies of rocky intertidal zones reveal that intense competition for space leads to competitive exclusion of inferior species (Connell 1961). The best competitor is rewarded with attachment space and a spatial monopoly (Connell 1961). Superior competitors can cause morality by crushing, prying, and pushing other organisms off through direct physical interference (Roughgarden et al. 1988).

31 My research focused on interference competition between C. virginica and a native

and an invasive barnacle in Mosquito Lagoon, FL (Fig. 1). This research is important as

competition is among the most important organizing factors within marine benthic

communities (Gurevitch et al. 2000). Much needed restoration efforts for the declining

eastern oyster, Crassostrea virginica have been masked by fouling organisms completely

covering submerged surfaces within days (Fig. 22). A better understanding of the

competitive interactions among sessile organisms that share limited resources gives insight

into the ecological conditions necessary for establishment and survival of oyster populations.

Only a limited number of studies have been conducted with Crassostrea virginica and

large densities of competing sessile species (Osman et al. 1989; Zajac et al. 1989; Osman et

al. 1990). Osman et al. (1989) found limited growth and mortality of newly settled spat to be

associated with the presence of other sessile invertebrates, including barnacles, ascidians and

bryozoans. It was later found that there were ontogenic changes in the relationships among

C. virginica and these interacting species (Osman et al. 1990). Accordingly, C. virginica’s susceptibility to certain competitors changed with the age of the oyster; for example a species that is initially a predator on oyster larvae may become a spatial competitor in a later life-stage (Osman et al. 1990). Zajac et al. (1989) found that even when competitors such as

Botrylloides sp., do not kill oysters, these competitors reduced the growth rate of Crassostrea

vriginica. From these examples it is evident that competition for attachment space and food

have a strong negative effect on settlement, growth and the survival of C. virginica (Jackson

1979; Buss 1981; Osman et al. 1989; Zajac et al. 1989; Osman et al. 1990).

The dominant competitors of oysters within Mosquito Lagoon are Balanus spp. The native barnacle, Balanus eburneus, and the invasive barnacle, Balanus amphitrite, have been observed in dense sets on oyster shells deployed within the IRL (Fig. 23). At certain times of

32 the year in Mosquito Lagoon, fouling organisms can completely cover unprotected surfaces

within days. Over 300 barnacles have been documented attached to one side of a single

disarticuleated oyster shell, (8 x 15 cm) within the IRL (Fig. 22).

Within the Mosquito Lagoon, Balanus spp. spawn year-round (Mook 1987). Peak

settlement occurs from July to November (Mook 1976). The native, acorn barnacle is the

ivory barnacle Balanus eburneus. Balanus eburneus is a large white concial barnacle, which grows

to 25 mm in both height and width (Mook 1980) (Fig. 23). This is the most prevalent

barnacle attached to oyster shells (Mook 1976; Boudreaux unpublished data). Its native range extends from Maine to South America (Mook 1980). The invasive acorn barnacle

Balanus amphitrite also inhabits Mosquito Lagoon (Zullo 1963). Although common, B.

amphitrite is usually not present in dense sets (Mook 1976). Balanus amphitrite is easily

distinguished by its fine gray to purple vertical stripes. It grows up to 9 mm in width and

originates from the Southwestern Pacific and Indian Oceans (Zullo 1963). Its Atlantic range

now extends from Cape Cod to the Caribbean. Its spread has been attributed to ship hull

fouling (Zullo 1963). It is a relatively recent invader to Mosquito Lagoon (within the past

100 years) ( J. Carlton pers comm.). It is important to include this invasive species in studies

of competitive interactions as it may be significantly altering the previous competitive

equilibrium that existed between the native barnacle, Balanus eburneus, and oyster populations.

Nothing is known about the interspecific competition of the oyster populations

within Mosquito Lagoon. Competitors of C. virginica have shown the potential to

significantly impede population size of C. virginica and may hinder the success of restoration.

Once the interactions of these competitors are determined, efforts to reduce these

competitive interactions will act to increase juvenile oyster success. This research will

ultimately aid in establishing the protocols needed to determine the optimal restoration

33 techniques of Crassostrea virgininca within Mosquito Lagoon. In particular, the timing of deployments for restoration should occur when barnacle impact is minimal. It will also answer important questions regarding recent oyster mortalities. All findings will protect and restore the functional integrity of Mosquito Lagoon, by helping to restore and maintain essential oyster habitat.

Methods

Competitor Effects on Oyster Settlement

To determine if Balanus spp. affect settlement of C. virginica, laboratory experiments were conducted. Densities of Balanus eburneus, B. amphitrite, and a combination of the two were manipulated on disarticulated oyster shells from Mosquito Lagoon. Densities of each treatment included: no barnacles (control), low (<25% barnacle cover), medium (25-50%), and high coverage (>50).

I obtained hatchery-reared competent Crassostrea virginica larvae from Middle Pensiula

AquaCulture in North, VA. and from Harbor Branch Oceanographic Institute in Ft. Pierce,

FL. Replicate laboratory settlement trials with cultured oyster larvae were run in both still water and flow (5 cm/s) using all density and species treatments in each flow treatment (Fig.

24). The dates of the experiments were August 2004 (still water) and May 2005 (flowing water).

Larvae were refrigerated prior to the experiments. One hour before each experiment, cyprids or oyster larvae were brought up to 24° C by placing them in filtered lagoon water

(Tables 17,18). Over the course of 60 minutes, oyster larvae and cyprids were repeatedly observed under a dissecting microscope (2.5 X) to determine larval activity. At least 50% of

34 the observed oysters were swimming or crawling on the bottom of the observation chamber

before experimentation began (Tamburri et al. 1996; Wall 2004). For all trials, larvae were suspended in a beaker and slowly poured from the beaker (over 5-10 sec) into the container

4 cm above the bottom (Tamburri et al. 1996; Wall 2004). Within each experimental date the order of trials was randomly selected to reduce any settlement differences in larval cohorts and relative larvae age in the experiments.

In all trials, larvae were allowed to settle on shell treatments for one hour with three replicate trials run on each date. All shell treatments were present at the same time. Three replicates of each shell treatment were present during each still water run for a total of 45 shells. Eight of each shell treatment were present in each flow trials for a total of 120 shells.

The still water treatments took place in a plastic tub (Sterilite Clearview 63 L, Model number

1753; 55 X 37 X 30 cm). Flow water treatments took place in a recirculating raceway flume.

Water volume and larvae were volume normalized for the containers. In still water, 20.0

liters of filtered lagoon water was added with 3,971 ± 914 oyster larvae. The flume used for

flowing water treatments was 20-cm wide, consisting of two semicircular ends (20-cm radius

at inner walls) and two straight sections, 120-cm long (Tamburri et al. 1996). Flow was

generated through the use of a motor-driven paddle wheel. Experiments were conducted at a

flow rate of 5 cm/s. The flow rate is the average mainstream flow rate found within

Mosquito Lagoon on a calm day (L.Walters, pers. com.). For both experiments, lagoon water

was obtained from the waters adjacent to the research station (salinity 30-33 ppt; 24-28ºC).

The water was filtered with a 25-micron mesh bag filter (Aquatic Eco-systems, Model

#N1025) to remove sediment, organisms, and debris. New filtered lagoon water was used in

each trial. Water volume was normalized for the container. The depth of water in both the

flume and the still water tub was 10 cm. The duration of each trial was 1 hour. Settlement of

35 oyster larvae was counted under a dissecting microscope (2.5x). The number of oyster larvae

added to the flow tank was 11,983 ± 689.

Competitor Effects on Oyster Growth and Survivorship

To determine if and how Balanus spp. affect the growth and survivorship of C. virginica, field experiments were conducted. Densities of Balanus eburneus and a combination treatment of Balanus eburneus and Balanus amphitrite were manipulated on oyster shells with 7- day old oyster spat from Mosquito Lagoon. Densities of each treatment included: 1) no barnacles (control), 2) low (< 25%) coverage of barnacles, 3) medium (25%-50%), and 4) high coverage (>50%). At the start of the trail all oyster were at maximum distance of 2 mm away from the nearest barnacle to ensure contact during the course of the study. All shell treatments were then deployed in Mosquito Lagoon waters in enclosures (1 m x 0.25 m) made of Vexar mesh (mesh size 0.25 cm) (Fig. 25). Each enclosure was constructed to compartmentalize and separate each individual shell and to prevent predation and minimize physical contact between shells that could induce oyster mortality (Fig. 25). Enclosures were deployed on two oyster reefs separated by 0.25 km within Mosquito Lagoon (6 enclosures/reef). Three shells of each treatment were present per enclosure for a total of 316 oyster shells per trial date. Two trials were run, one in October/November 2004, and one in

July/August 2005. Competitive interactions between the oysters and barnacles were measured by contrasting the growth and survivorship of juvenile Crassostrea virginica from the different shell treatments to the controls with no barnacles. Growth and survivorship were monitored every 3 days for 4 weeks by use of transparency film. I analyzed these data using

SPSS and conducted a two-way ANOVA testing whether C. virginica growth varied as a

36 function of Balanus spp. shell treatments (zero, low, medium, high) or trial date.

Comparisons of survival rates over the four weeks were measured using Wilcoxon statistics.

Results

Settlement of larvae of C. virginica was not consistently affected by the presence of

either species of Balanus. the still water treatment was significantly different from the flowing

water treatment (p=0.040) (Fig. 26; Table 12). In still water, there were no differences in

settlement among any of the densities or species treatments. In the second flowing water,

only the control received significantly higher settlement than all other treatments (p=0.04)

(Fig. 26).

Growth of C. virginica was significantly affected by the presence of Balanus spp.

Control treatments (without any barnacles) had a significantly higher mean growth over the

course of the experiments (9.9 + 0.3 mm; Mean + S.E.) than all other treatments (5.7 + 0.6 mm) (p< 0.001) (Fig. 27; Table 13). Growth was not significantly different among any of the densities or species treatments besides the control although the interaction was significant

(Fig 27; Table 13). Growth rates did not vary between the trial dates (p=0.38).

Survival rates of Crassostrea virginica were significantly different over the course of 4 weeks in the two trials (p<0.001)(Figs. 28, 29). Trial one showed significant differences only between the control and all other treatments (p<0.001)(Table 14). In trial two, the control,

B. eburneus low and B. eburneus medium treatments had significantly higher survival than all other treatments (p=0.006)(Fig. 29; Table 15).

37 Discussion

Many marine sessile organisms choose habitat locations which will increase their

post-settlement success (e.g. Scheltema 1974; Grosberg 1981; Branch 1984, Young 1990;

Dalby and Young 1993). Selection to avoid opponents can drive genetic differences on

microevolutionary timescales (Sotka 2005). In the case of the crab Libinia dubia and the

amphipod Amphitote longimana, there is experimental evidence that habitat choices mediate

susceptibility to predators (Sotka 2005). Additionally, sessile bryozoan larvae tend to settle to

avoid competition with algae (Ryland 1977). Does C. virginica minimize post-settlement mortality due to the spatial competition the same way? It may be beneficial for C. virginica to avoid settling in locations with high Balanus spp. densities since they are not able to survive or grow well in these locations. This study found that the settlement of C. virginica was not consistently affected by the presence of Balanus spp. Trial one in still water yielded no significant difference in settlement among any density or barnacle species treatment. These oysters settled in similar numbers on the disarticulated oyster shells with no barnacles as they did on shells with high barnacle density. Additionally there were no consistent patterns with the two separate barnacle species. Trial two however, which occurred in flowing water of

5cm/s did show greater settlement on the biolfilmed shells with no barnacles (Fig. 26). This trend is important as it shows that in normal conditions in the lagoon the oysters are able to preferentially settle on substrate in which they have a higher probability of survival.

Despite the abundance of literature on the ecology of the oyster C. virginica, few field experiments have investigated the actual competitive abilities of this species (e.g. Bros 1987;

Bushek 1998; Osman et al. 1989; Zajac et al. 1989; Osman et al. 1990; Dalby and Young

1993). In all cases, the growth and survivorship of the eastern oyster was negatively effected by the presence of other sessile species. In Mosquito Lagoon, growth of juvenile C. virginica

38 was shown to be reduced in the presence of barnacles, no matter the density or species

(native vs. invasive) in contact with the juvenile oyster. Trial date did not alter the outcome.

Growth rates of juvenile oysters in Mosquito Lagoon were similar in the summer (July-

August 2005) and the late fall (October-November 2004).

Survivorship of C. virginica was greatest when there were no barnacles present, and then decreased as the density of barnacles per shell increased on the treatment (Figs. 27, 28).

The specific barnacle species did not change the survival rates over the course of the study.

This study indicates that the specific species of barnacle in contact with the oyster does not affect the outcome. The mere presence of any barnacle species decreases its chances of survival.

Research has been conducted comparing the total number of barnacles on extant oyster shells to historic midden shells (see Chapter 1). The introduction of B. amphitrite to

Mosquito Lagoon, FL within the past 100 years has increased the total number of barnacles per oyster shell by 5-fold. The mere presence of barnacles, independent of barnacle species, has been shown to decrease juvenile oyster survival and growth; the addition of B. amphititre has greatly increased the overall spatial competition between oysters and barnacles.

The structure of populations of marine organisms can be determined by processes affecting the arrival and early survival of new individuals (Roegner 1991). Understanding the role that Balanus spp. competitors play in affecting the settlement, growth and survival of juvenile C. virginica is critical for a better understanding of the forces shaping these fragile oyster populations in Mosquito Lagoon, FL.

39

CHAPTER THREE:

EFFECT OF FLOW AND SEDIMENT ON THE SETTLEMENT OF OYSTER

AND BARNACLE LARVAE

Introduction

Although the ecological importance of oysters has been known for at least a century,

they have historically been treated as a species to exploit rather than conserve (Micheli &

Peterson 1999). Recently, concern for this species has spread throughout the entire east

coast of the United States (Micheli & Peterson 1999). Increasing anthropogenic,

disturbances along the coast have led to substantial losses in eastern oyster populations

(Coen et al. 1999). Threats include overfishing, disease, and habitat degradation (Coen et al.

1999). The economic and ecological importance of this species therefore calls for efficient

approaches to the conservation and management of wild populations. It is vital to protect

and restore these refuges of biodiversity. Mosquito Lagoon, Florida (Fig. 1) is one of the few

areas with relatively undisturbed intertidal reefs of the eastern oyster Crassostrea virginica in the

United States and is the southernmost extent of this species distribution along the USA

Atlantic coast. The preservation of these unique populations is vital.

Populations of C. virginica in Mosquito Lagoon have recently undergone significant die-offs and are a subject of major concern by resource managers. Some reefs of C. virginica have declined by 50% in total area from 1943 to 1995 (Grizzle et al. 2002). Restoration efforts have begun within Mosquito Lagoon and are focusing on reconstructing the three- dimensional reef habitats (L. Walters pers comm.). These efforts hope to re-establish the

40 ecological role of what are now merely footprints of former reefs. Before the most effective

protocol can be established, a better understanding of the ecology of these reefs is needed.

Aerial imagery and field studies have documented that reefs along major boating

channels have dead zones of disarticulated shells along the seaward edges of the oyster reefs

(Fig. 2) (Grizzle & Castagna 1995; Grizzle et al. 2002). It has been documented in an hour

approximately 50 boats pass by single oyster reef in the IRL. In Mosquito Lagoon, this

intense year-round boating has been held responsible for these dead margins arising and the

increases in oyster mortality (Wall et al. in press). More specifically boating is changing the

hydrographic conditions in Mosquito Lagoon, both increasing sediment loads and water

motion on oyster reefs (Wall et al. in press).

Now we need to understand and quantify the negative effects of increased water

motion and high levels of sedimentation on larval settlement to determine which

mechanisms are causing the oyster reef declines (Grizzle et al. 2002). The potential negative

effects on larval forms of flow and sediment levels associated with boating activity on needs

to be determined.

The purple striped barnacle, Balanus amphitrite, invaded Mosquito Lagoon

approximately 100 years ago via ballast water transport (J.Carlton pers. comm.). It now

competes for space with juvenile C. virginica. Over 300 barnacles, including the native Balanus

eburneus and the invasive, Balanus amphitrite, have been counted on a single oyster shell (Wall

2004). This high number of spatial competitors may be decreasing the success of Crassostrea

virginica. If B. amphitrite can settle in a wider variety of flow rates and sediment conditions

than Crassostrea virginica, then it may have a competitive advantage over oysters in this space-

limited habitat.

41 Previous research showed that C. virginica settlement decreased in response to 5 cm/s flow and increased sediment loads (0, 8, 16 g/ml) (Wall 2004). I expanded on this research to determine how C. virginica responds to unnaturally high flow from boating (10 cm/sec).

Additionally, I quantified the effects of water motion (0, 5 cm/s, 10 cm/s) and sediment loads (0, 8, 16 g/ml) on the recruitment of an important, relatively new oyster competitor in the Indian River Lagoon system, the barnacle Balanus amphitrite.

Methods

Study Site

Research was conducted in Mosquito Lagoon, within Canaveral National Seashore

(Fig. 1). The Lagoon system is a series of three distinct, but connected, estuaries which extend 251 kilometers (156 miles) from Ponce de Leon Inlet to Jupiter Inlet on the east coast of central Florida. The University of Central Florida research facility, Fellers House

Field Station (28° 54’ N, 80° 49’ W), is located within the bounds of the National Park.

The average depth of the Lagoon is less than 1 meter in most areas and the current is primarily wind-driven (Walters et al. 2001). Annual salinity ranges between 18 and 45 ppt, depending on rainfall (Grizzle 1990; Walters et al. 2001; Boudreaux unpublished data). Most of the lagoon is a complex system of shallow and open water areas with nearly 100 mangrove (Rhizophora mangle and Avicennia germinans) dominated islands (Walters et al. 2001).

Intertidal oyster reefs are found throughout this region.

Laboratory experiments

Laboratory experiments were performed with larvae of the oyster Crassostrea virginica and the barnacle Balanus amphitrite to determine which of these competitors were better able

42 to withstand increasing flow and sediment conditions. The methods and sediment loads are taken from Wall (2004). Thirty six separate trials were conducted. For C. virginica, three replicates were run for each sediment load at 10 cm/s flow. Three replicates were run for each sediment load at 0 cm/s, 5 cm/s, and 10 cm/s for the barnacle larvae. All trials were run at three sediment levels: no sediment, low sediment (8 g/ml), and high sediment (16 g/ml); loads for each of the specified flow rates (Table 16). The experiments with the oyster larvae took place in May 2005 and the experiments with the barnacle larvae took place in

October 2004 and May 2005 (Figs. 19, 20).

Cyprid larvae of Balanus amphitrite were obtained from Dr. Dan Rittschof at Duke

University Marine Lab and shipped on ice via overnight courier to the University of Central

Florida. Larvae of C. virgininca were obtained from Dr. John Scarpa at Harbor Branch

Oceanographic Institute with permission from William Arnold of Florida Fish and Wildlife

Institute. All larvae were kept refrigerated at 7° C until experiments were run (<48 hrs after delivery). Within each experimental date the order of trials was randomly selected to reduce any settlement differences in larval cohorts and relative larvae age in the experiments.

One hour before each experiment, cyprids or oyster larvae were brought up to 24° C by placing them in filtered lagoon water (Tables 17,18). Over the course of 60 minutes, oyster larvae and cyprids were repeatedly observed under a dissecting microscope (2.5 X) to determine larval activity. At least 50% of the observed oyster and barnacle cyprids had to be swimming or crawling on the bottom of the observation chamber before experimentation began (Tamburri et al. 1996; Wall 2004). For all trials, cyprids were suspended in a beaker and slowly poured from the beaker (over 5-10 sec) into the container 4 cm above the bottom

(Tamburri et al. 1996; Wall 2004).

43 Sediment was collected from the seaward regions of nearby oyster reefs of Mosquito

Lagoon immediately before each trial (Wall 2004). Sediment loads were normalized by total volume of water in the tank and tub (Fig. 17). Three replicate trials were conducted with each of the three levels of sediment for both the still-water and flowing water experiments.

Trials had no sediment, low sediment loads (8 g/ml wet weight), or high sediment loads, (16 g/ml wet weight).

Disarticulated oyster shells were used as substrate in all settlement trials. Prior to each set of still and flow trials, 4500 disarticulated oyster shells were cleaned to remove all macroflora or macrofauna and placed into the Lagoon for 6 days to establish a new natural biological film. Immediately prior to use, shells were visually inspected, and any with attached macroflora or macrofauna were not used in the experiment. Lagoon water, obtained from the waters adjacent to the research station (salinity 30-33‰; 24-28 ºC), was filtered with a 25 micron mesh bag filter (Aquatic Eco-Systems, Model number N1025) to remove sediment, organisms, and debris. New filtered lagoon water was used for each run.

The duration of the oyster trials were one hour. The barnacle trials were conducted over 24 hours in the dark.

Still-water experiments were conducted at the same time as the flowing-water experiments in a plastic tub (Sterilite Clearview 63 L, Model number 1753; 55 X 37 X 30 cm). Water volume and larvae used were normalized. For each trial, 20.0 liters of filtered lagoon water were added. The depth of water was 10 cm. Seventy oyster shells, half of which had the inside of the valve facing up and half had the outside of the valve facing up, covered the bottom of the container (Fig. 24). The volume normalized number of oyster larvae added was 4,215 ± 451. Additionally 3,660 ± 123 cyprids were added to the still water tub for the barnacle trials.

44 Flowing-water experiments were conducted in a recirculating raceway flume. The

flume was 20-cm wide, consisting of two semicircular ends (20-cm radius at inner walls) and

two straight sections, 120-cm long (Tamburri et al. 1996). Water flow was generated through the use of a motor-driven paddle wheel. To reduce across-stream fluid motion, polycarbonate sheeting was place parallel to the curved flume walls upstream of the working area to straighten the flow. One hundred and forty oyster shells, half of which had the inside of the valve facing up and half had the outside of the valve facing up, covered the bottom of

the tank. Eighty liters of filtered lagoon water was added for a depth of 10 cm. the number

of oyster larvae added were 12,833 ± 4585; cyprids 6,120 ± 697.

After each trial, shells were gently removed from the water and observed under a

dissecting microscope. Settlers were counted as individuals attached to shells. After each trial, the plastic container and the flume were rinsed with freshwater and dried to remove any remaining oyster larvae, barnacle cyprids and sediment.

Analysis I used SPSS to run separate 2-way analysis of variance (ANOVA) for each invertebrate. All factors were fixed: flow rate (flow: 5 cm/s or still-water: 0 cm/s) and sediment load (high, low, or no sediment). Prior to running the ANOVAs, I tested for normality and heterogeneity, using Levene’s F and Kolmagorov-Smirnov tests. If a significant difference was found, a Bonferroni’s pairwise comparison was used to determine differences between treatments at a significance difference level at α = 0.05 with 95% confidence intervals.

45 Results

Data for oyster trials at 10 cm/s were combined with previous data from Wall (2004)

who examined oyster settlement at 5 cm/s and in still water. Settlement of larvae of the

oyster C. virginica differed significantly due to both flow rate and sediment load (Fig. 30;

Table 19). The interaction of flow rate and sediment load was significant (p< 0.001). This

interaction is driven by low settlement in the no sediment 10 cm/s treatment. Oyster larval

settlement was significantly lower in flowing-water (5 cm/s; 10 cm/s) compared to still-

water (p = 0.0020) (Fig 30). There was no difference in settlement between 5cm/s and

10cm/s treatments (Fig. 30). Larval settlement in low sediment loads was similar to

settlement in no sediment trials. Conversely, settlement of barnacles cyprids did not differ

significantly due to either flow rate (p=0.598) or sediment load (p=0.244) (Fig. 31; Table 20).

Discussion

This study was the first step in identifying and quantifying the potential negative

effects of increased sediment loads and flow rates in Mosquito Lagoon. Knowledge of larval

settlement in site-specific conditions is critical to understanding regional population

dynamics (Coen and Luckenbach 2000) and will aid in the development of management and

restoration plans for this unique intertidal system.

Studies support the notion that sediment resuspension and flow rate may decrease oyster recruitment (Shelbourne 1957; Seliger et al. 1982; Lenihan 1999; Wall 2004).

Crassostrea virginica was affected by both sediment and loads even at the low levels we tested

(Fig. 30). It was found that settlement of oyster larvae decreased with increasing sediment

loads. The negative effects of sedimentation on oyster reef have also been noted in many other field studies (MacKenzie 1977; Gunter 1979; MacKenzie 1983; MacKenzie 1996a,

46 1996b; Bartol et al. 1999). These high levels of sedimentation reduced oyster larval

settlement by altering the surface topography. Altering surface topography influences larval

settlement (Walters 1992) and sediment in constant motion may cause mortality of settling cyprids by abrasion (MacKenzie 1996b). Often, changes in the flow rates can explain differences in sedimentation rates seen on oyster reefs (Lenihan 1999). Wave action (i.e. increased flow rate) may lead to an accumulation of sediment eventually smothering oysters and high turbidity may decrease larval set (Kennedy & Sandford 1999; Bartol et al. 1999).

Flow may also influence the delivery of larvae to the substrate and the maintenance of position during and after settlement (Nowell and Jumars 1984). Additionally, Balanus spp. resilience to increased sediment loads or and flow is supported by literature. Balanus spp. have a very high tolerance to sediment and flow rate (1 m/s in the field) (Mangum et al.

1972).

In these experiments the methods used have some limitations for the interpretation of the data. The flow tank used produced an average mainstream flow rate of 5 cm/s or 10 cm/s. Every location within the tank was not experiencing these flows. Additionally, a flow rate of 5 cm/s in the flow tank setting should be compared very carefully to 5 cm/s flow in the field. The dynamics affecting flow in the tank at a water depth of 10 cm is undoubtedly different from the same lagoonal flow (water depth 1 m). The sheer stress and mainstream velocities experienced by individual larvae will be much greater in the flow tank than in the field.

Flow and sediment reduced the larval settlement of C. virginica. The ability of barnacles to settle efficiently in 5 cm/sec (natural lagoon flow) and at 10 cm/sec (increased flow due to boat wakes or storms) and not be affected by sediment gives them a competitive advantage. The intense recreational boating activities in Mosquito Lagoon are increasing the

47 flow rate to unnaturally high conditions (>5 cm/s) and causing harmful sediment resuspension. Thus, continuous boat traffic during settlement will favor recruitment of the invasive barnacle Balanus amphitrite over the native oyster Crassostrea virginica.

48

APPENDIX A- FIGURES

49

Figure 1- Indian River Lagoon system (Smithsonian 2001)

50

Figure 2- Oyster reefs Left: An impacted oyster reef in Mosquito Lagoon with uncharacteristic dead margin on the seaward edge of the reef at high tide. Right: a pristine (unimpacted) reef in Mosquito Lagoon at low tide.

51

Figure 3- Internal diagram of Crassostrea virginica (Ashbaugh 1951).

52

Figure 4- Life-cycle of Crassostrea virginica (Wallace 2001).

53

Figure 5- Lift nets study sites in Mosquito Lagoon, Florida

54

Figure 6- Submerged lift net (1m2)

Figure 7- Lift net retrieval

55 5

S.E. 4.5 4 3.5 3 2.5 2 1.5 1 0.5 0

5 04 -04 04 0 -05 -05 n- t- c-04 b- n Mean species richness + richness species Mean u e e pr J Aug Oc D F A Ju

Figure 8- Mean richness (total number of species) ± S.E. per net per month *September data missing due to 2004 Hurricane Jeanne.

60 50 40 30 20 10 S.E. 0

5 04 -04 -04 0 05 -05 t-04 c - n un- ug J A Oc De Feb- Apr Ju Mean percent dominance + percent dominance Mean Sampling Period

Figure 9- Mean percent dominance (± S.E.) of all lift nets per month *September data missing due to 2004 Hurricane season

56 60 50 40 spp. S.E. 30 20

Balanus 10 0 abundance + abundance Mean Mean 04 05 04 -04 04 -05 Jun- Aug- Oct Dec- Feb- Apr Jun-05 Sampling Period

Figure 10-Mean Balanus spp. abundance (± S.E.) per net *September data missing due to 2004 Hurricane Jeanne.

5

4

3

2

1 Mean Richness Mean 0 Live Oysters Disarticulated Shells Settlement substrate

Figure 11-Mean richness (± S.E.) on live oysters compared to disarticulated shells

57 60

50

40

30

20

10 Mean Dominance 0 Live Oysters Disarticulated Shells Settlement substrate

Figure 12- Mean percent dominance (± S.E.) on live oysters compared to disarticulated shells

35

30 25 spp.

S.E. 20 15 10 Balanus 5 0 Mean abundance + Live Oysters Disarticulated Shells Settlement Substrate

Figure 13- Mean Balanus spp. abundance (± S.E.) on live oysters compared to disarticulated shells

58

2 1.5 1 0.5

Mean Diversity 0

4 5 5 05 -0 n-0 -04 b-0 - u ug-04 ct e pr un J A O Dec-04 F A J Sampling Period

Figure 14- Mean diversity (± S.E.) of lift nets per month *September data missing due to 2004 Hurricane season

1.5

1

0.5 Mean Diversity 0 Live Oysters Disarticulated Shells Substrate type

Figure 15-Diversity on live oysters compared to disarticulated shells

59 500 400 300 200 100 0

Mean sediment load (g) rch June mber a May ugust M A January Nove

Figure 16-Mean sediment (± S.E.) load per month from June 2004- June 2005

120 100 80 60 40 20

Sediment 0

r y y ne e r ch u a r Ma J nu a emb a M August v J

Silt/Clay Percentage of Total No

Figure 17- Mean silt/clay percentage (± S.E.) per month from June 2004-June 2005

60 40 35 30 25 20 15 10 5 0 Mean salinity (ppt) Mean salinity 4 4 5 0 05 n-0 - r-0 u p un-05 J Aug-04 Oct- Dec-04 Feb A J

Figure 18- Mean salinities (± S.E.) of Mosquito Lagoon on lift net sampling dates

35 30

S.E. S.E. 25 20 15 10 5 0 Degrees celsius

Mean temperature + temperature Mean -04 05 ec-04 Jun-04 Aug Oct-04 D Feb-05 Apr- Jun-05

Figure 19- Mean monthly temperatures (± S.E.) from lift net sampling dates in Mosquito Lagoon

61 X Extant Comparison Site

Lift net Sites

Extant Comparison Site X

Extant Comparison Site X

Figure 20- Historic shell midden sites (=archeological sites) and nearby extant oyster reef sites

62 9 8 7 Extant 6 5 Historic 4 3 2 1

Maximum dimension (cm) 0 Length Width

Figure 21- Comparison of size of midden to extant oyster shells

Figure 22- Example of an oyster shell covered with barnacles.

63 A B

Figure 23- Balanus spp. of Mosquito Lagoon. A: B. amphitrite and B: B. eburneus.

A B

Figure 24- Oyster settlement experimental set-up. A: Recirculating flow tank. B: Still water tub

64

Figure 25-Field enclosure for oyster growth and survivorship experiment

65 350 300 250 200 150

treatment 100 50 0

Mean oyster settlement per shell per per shell per settlement oyster Mean Still Water Flow 5 cm/s

No Balanus spp. B. amphitrite low B. amphitrite medium B. amphitrite high B. eburneus low B. eburneus medium B. eburneus high Combination low Combination medium Combination high Figure 26-Oyster settlement (± S.E.) on barnacle densities and species treatments.

Densities of Balanus eburneus, Balanus amphitrite, and a combination of Balanus eburneus and Balanus amphitrite were manipulated on disarticulated oyster shells from Mosquito Lagoon. Densities of each treatment included: no barnacles (control), low (<25% barnacle cover), medium (25-50%), and high coverage (>50 %). Trial one was conducted in still water; Trial two flow 5 cm/s.

66

12

10

8

6 weeks 4

2

Mean spat growth (mm) over 4 over (mm) growth spat Mean 0 Control Zero Barnacles Balanus eburneus low Balanus eburneus mediumTreatmentsBalanus eburneus high Combination low Combination medium Combination high

Figure 27-Mean growth (± S.E.) over 4 weeks. Densities of Balanus eburneus, and a combination of Balanus eburneus and Balanus amphitrite were manipulated on disarticulated oyster shells from Mosquito Lagoon. Densities of each treatment included: no barnacles (control), low (<25% barnacle cover), medium (25-50%), and high coverage (>50%).

67

1 l 0.8 0.6 0.4

Cumulative Surviva Cumulative 0.2 0 1 4 7 1 4 8 2 5 8 y y y 1 1 1 2 2 2 y y ay y y y Da Da Da Da Da D Da Da Da

Control Zero Barnacles Balanus eburneus low Balanus eburneus medium Balanus eburneus high Combination low Combination medium Combination high

Figure 28- Cumulative survival of C. virginica over 4 weeks: Trial 1 (October- November 2004)

68

1

0.8

0.6

0.4

0.2 Cumulative Survival 0

1 1 1 14 18 22 25 y Day Day 4 Day 7 ay D Da Day Day Day Day 28 Control Zero Barnacles Balanus eburneus low Balanus eburneus medium Balanus eburneus high Combination low Combination medium Combination high

Figure 29- Cumulative survival of C. virginica over 4 weeks; Trial 2 (July- August 2005)

69 Settlement of Crassostrea virginica

300 240 180 120 S.E.) 60 ( + 0 No No No Mean settlement of larvae Low Low Low High High High Sediment Sediment Sediment Sediment Sediment Sediment Sediment Sediment Sediment Still Water Flow: 5 cm/s Flow: 10 cm/s

Figure 30-Sediment and flow rate effect on the settlement of the oyster Crassostrea virginica. Sediment loads were 0 g/ml; 8 g/ml; 16 g/ml. Means of 3 replicates for each different treatment

Settlement of Balanus amphitrite

80

40 S.E.) 0 (+ No No No Low Low Low High High High Sediment Sediment Sediment Sediment Sediment Sediment Sediment Mean settlementMean of cyprids Sediment Sediment Sediment Still Water Flow: 5 cm/s Flow: 10cm/s

Figure 31-Sediment and flow rate effect on settlement of the barnacle Balanus amphitrite. Sediment loads were 0 g/ml; 8 g/ml; 16 g/ml. Means of 3 replicates for each different treatment

70

APPENDIX B- TABLES

71

Table 1- Lift net deployment and collection dates Deployed Collected June 13, 2005 July 13, 2005 July 13, 2005 August 4, 2005 August 4, 2004 September 1, 2004 October 1, 2004 November 6, 2004 November 6, 2004 December 12, 2004 December 12, 2004 January 8, 2005 January 8, 2005 February 5, 2005 February 5, 2005 March 5, 2005 March 5, 2005 April 2, 2005 April 2, 2005 April 30, 2005 April 30, 2005 May 28, 2005 May 28, 2005 June 25, 2005

72 Table 2- Total numbers of sessile species recruited to shells in lift nets

Common 6/04 7/04 8/04 10/04 11/04 12/04 1/05 2/05 3/05 4/05 5/05 6/05 Phyla Species Name Total Balanus Ivory 2070 2447 1380 720 430 210 145 87 132 99 251 811 Arthropoda eburneus Barnacle 8782 *Purple 438 461 450 46 16 13 9 1 5 6 8 71 Balanus Striped amphitrite Barnacle* 1524 *Tube 658 491 760 250 134 111 66 33 65 55 42 177 Annelida Hydroides spp. Worms* 2842 Feather 2 1 9 6 4 6 7 0 0 1 9 1 Duster Sabella spp. Worm 46 Sea 0 0 0 0 0 0 2 0 0 0 0 0 Cnidaria Aiptasia pallida anemone 2 Haliplanella *Striped 0 0 0 0 0 0 0 0 1 1 0 0 luciae Anemone* 2 Anomia 186 178 184 151 93 83 41 4 43 45 80 32 Mollusca simplex Jingle Shell 1120 Eastern 287 154 207 149 69 59 31 4 25 44 73 76 Crepidula Slipper astrasolea Shell 1178 Atlantic 12 0 3 4 4 0 9 0 1 1 0 6 Crepidula Slipper fornicata Shell 40 Diodora Keyhole 1 1 1 0 0 0 0 0 0 0 0 00 cayensis Limpet 3 Atrina rigida Pen Shell 1 0 0 0 0 0 1 0 0 0 0 0 0 Tangelus 0 0 0 0 0 0 0 1 0 0 0 0 divisus Jacknife 1

73 Clam Brachidonetes Scorched 0 0 2 0 0 1 0 1 0 0 0 0 exuctus Mussel 4 Geukensia Ribbed 21 20 19 7 6 4 7 0 2 4 19 18 demissa Mussel 128 Mytella *Charru 0 0 0 3 0 0 0 0 0 0 0 0 charruana Mussel* 3 Lithophaga Mahogany 0 0 0 0 0 0 0 0 0 0 0 1 bisulcata Date 1 Hymeniacidon Sun 2 2 1 0 11 0 3 0 1 2 4 2 Porifera heliophila Sponge 28 Black 1 0 0 0 0 5 0 0 0 0 3 57 Halichondria Volcano melandocia Sponge 66 Boring 0 1 0 0 0 0 9 1 2 0 0 0 Ectoprocta Cliona spp Sponge 13 *Common 1 1 0 0 2 0 16 0 18 82 48 27 Bugula neritina Bryozoan* 195 Hippoporina Lacy 0 0 1 28 11 0 0 0 0 0 0 0 verrilli Bryozoan 40 Zoobotryon Spaghetti 0 0 0 0 0 0 0 0 0 0 0 2 verticillatum Bryozoan 2 Perophera Encrusting 0 2 0 1 5 3 3 0 2 0 0 0 Chordata viridis Ascidian 16 Rough Sea 0 0 0 26 17 2 1 0 1 15 6 19 Styela plicata Squirt 87 *denotes invasive species

74

Table 3- ANOVA comparison of species richness in lift nets. The factors were month, reef type (pristine or dead margins), and shell type (live or disarticulated oyster).

Source df Mean Square F P value MONTH 11 85.401 34.576 0.000 REEFTYPE 1 64.201 4.625 0.098 SHELLTYPE * 6 104.321 42.236 0.000 REEFTYPE * MONTH Error 697

Table 4- ANOVA comparison of barnacle abundance in lift nets. The factors were month, reef type (pristine or dead margins), and shell type (live or disarticulated oyster).

Source df Mean Square F P value MONTH 11 16645.381 6.839 0.000 REEFTYPE 1 38984.450 2.238 0.209 SHELLTYPE * 6 29478.006 12.111 0.000 REEFTYPE Error 697

Table 5- ANOVA comparison of percent community dominance in lift nets. The factors were month, reef type (pristine or dead margins), and shell type (live or disarticulated oyster).

Source df Mean Square F P value MONTH 11 3121.876 4.010 0.000 REEFTYPE 1 1311.168 .619 0.475 SHELLTYPE * REEFTYPE 6 20374.641 26.170 0.000 Error 516 778.535

75

Table 6- ANOVA comparison of diversity found in lift nets. The factors were month, reef type (pristine or dead margins), and shell type (live or disarticulated oyster).

Source df Mean Square F P value MONTH 11 1.028 5.343 0.095 REEFTYPE 1 1.560 2.560 0.184 REEFTYPE * 6 .933 4.845 0.159 SHELLTYPE Error 516 .192

Table 7- ANOVA comparison of total sediment loads collected per month. The factors were month and reef type (pristine or dead margins).

Source Df Mean Square F P value REEFTYPE * 75 36911.908 .792 0.872 MONTH MONTH 11 52155.362 1.119 0.320 REEFTYPE 1 77711.307 1.668 0.198 Error 164 46590.791

Table 8- ANOVA comparison of silt/clay fraction of sediment loads collected per month. The factors were month and reef type (pristine or dead margins).

Source Df Mean Square F P value REEFTYPE 1 313.463 .449 0.504 MONTH 12 4044.778 5.794 0.000 REEFTYPE * 23 340.027 .487 0.745 MONTH Error 183 698.128

76 Table 9-Shell midden historic sites and closest extant oyster reefs in Mosquito Lagoon

Middens GPS Approx. collection date (Radiocarbon)-dating

Oyster Bay 28◦57’30.13N A.D. 250-940 80◦51’43.91W

Turtle Mound 28◦55’49.30N A.D. 750-1550 80◦49’37.48.91W

Seminole Rest 28◦52’35.76N A.D. 280-1420 80◦50’28.45W

Extant Reefs

Oyster Bay 28◦57’26.75 N 80◦51’43.97 W

Turtle Mound 28° 55.887 N 80°49.859 W

Seminole Rest 28° 53.217 N 80°51.198 W

77 Table 10- Separate one-way ANOVAs comparing midden versus extant oyster shell sizes (length and width) LENGTH df Mean Square F P value SHELL 543 313.463 .18 0.443 Error 183 588.128 20.55

WIDTH df Mean Square F P value SHELL 543 1162.463 52.48 0.789 Error 183 354.128 18.02

Table 11- ANOVA to compare settlement of barnacles on historic versus extant shells. Shell source = midden or extant shells. Site is the collection zone (Turtle Mound, Oyster Bay or Seminole Rest).

Source df Mean Square F P value SITE 2 45.606 7.891 0.000 SHELL SOURCE 1 221.048 38.249 0.000 SITE * SHELL 2 150.500 26.041 0.000 SOURCE Error 623 5.779

78

Table 12-Settlement of oysters on shells with varying barnacle density and species either in flow and still water Source df Mean Square F P value Flow 1 8780.852 4.538 0.046 Barnacle Treatment 9 6859.476 3.545 0.009 Flowt* Barnacle Treatment 9 9028.692 4.666 0.002 Error 20 1934.933

79 Table 13- ANOVA comparison of oyster growth at the end of four weeks Source df Mean Square F P value

Trial Date 1 6.875 0.746 0.389 Treatment 6 74.590 8.091 0.000 Trial * Treatment 6 42.001 4.556 0.000 Error 13 9.219

80

Table 14-Wilcoxon survivorship statistics- Comparison of treatments in Trial 1 Treatment Mean Score Control 38.3143*

B. eburneus low 2.4583

B. eburneus medium -28.833

B. eburneus high -42.4783

B. eburneus and B. amphitrite low -.13.800

B. eburneus and B. amphitrite medium -26.3478

B. eburneus and B. amphitrite high -5.0147

*denotes significantly different

81 Table 15- Wilcoxon survivorship statistics- Comparison of treatments in Trial 2 Treatment Mean Score Control 19.4776* B. eburneus low -1.000*

B. eburneus medium -2.8235*

B. eburneus high -46.2500

B. eburneus and B. amphitrite low -20.000

B. eburneus and B. amphitrite medium -39.000

B. eburneus and B. amphitrite high -106.000

*denotes significantly different

82

Table 16- Total amount of sediment used in trials Sediment Flow Treatment Sediment Load Treatment 1 = no sediment Flow tank 0 g (0 g/ ml) 2 = low sediment Flow tank 725.0 g (8 g/ ml) 3 = high sediment Flow tank 1450.0 g (16 g/ ml) 1 = no sediment Still water 0 g (0 g/ ml) 2 = low sediment Still water 145.0 g (8 g/ ml) 3 = high sediment Still water 290.0 g (16 g/ ml)

83 Table 17- Sediment and flow experiments: barnacles trials.

Trial Date Speed Sediment Temp. Salinity (cm s-1) load ºC (‰) 1 10/07/04 0 Low 26 30 2 10/07/04 5 Low 26 30

3 10/08/04 0 High 26 30 4 10/08/04 5 High 26 30 5 10/09/04 0 Zero 26 30

6 10/09/04 5 Zero 26 30 7 10/26/04 0 Zero 24 33 8 10/26/04 5 Zero 24 33

9 10/27/04 0 Low 24 33 10 10/27/04 5 Low 24 33 11 10/28/04 0 High 24 33 12 10/28/04 5 High 24 33 13 11/09/04 0 Zero 24 30 14 11/09/04 5 Zero 24 30 15 11/10/04 0 High 24 30 16 11/10/04 5 High 24 30 17 11/11/04 0 Low 24 30 18 11/11/04 5 Low 24 30 19 5/9/05 10 Low 24 34 20 5/10/05 10 High 24 34 21 5/11/05 10 Zero 24 34 22 6/7/05 10 Zero 24 34 23 6/8/05 10 High 24 34 24 6/9/05 10 Low 24 34 25 6/16/05 10 Low 24 34 26 6/17/05 10 High 24 34 27 6/18/05 10 Zero 24 34

84

Table 18- Sediment and flow experiments: oysters trials

Trial Date Speed Sediment Temp. Salinity

(cm s -1) load ºC (‰)

28 5/20/05 10 Low 26 30 29 5/20/05 10 High 26 30 30 5/20/05 10 Zero 26 30 31 5/20/05 10 Zero 24 33 32 5/21/05 10 Low 24 33 33 5/21/05 10 High 24 33 34 5/21/05 10 Zero 24 30 35 5/21/05 10 High 24 30 36 5/21/05 10 Low 24 30

85

Table 19-ANOVA comparison settlement of the oyster Crassostrea virginica in varying flow speeds and sediment loads Source df Mean Square F P value SEDIMENT 2 164786.17 180.23 < 0.0001* FLOW 2 23544.5 115.47 0.0020* SEDIMENT * FLOW 4 17470.5 11.48 < 0.0001* *Significant values

Table 20- ANOVA comparison settlement of the barnacle Balanus amphitrite at varying flow speeds and sediment loads Source df Mean Square F P value SEDIMENT 2 299.565 0.530 0.598 FLOW 2 863.370 1.526 0.244 SEDIMENT * FLOW 4 647.037 1.144 0.368

86 REFERENCES

Andrews, J.D. 1979. Pelecypoda Ostreidae. In a Reproduction of Marine Invertebrates, edited by .C. Glese

and Pearse, Academic press, New York, pp 293-341.

Andrews, J.D. 1991. The oyster fishery of eastern North American based on Crassostrea virginica. In

Estuarine and Marine Bivalve Mollusk, edited by W. Menzel, CRC Press, Boston, pp 11-23.

Ashbaugh, B.L. 1951. Things to Do in Conservation. Department of Research and Education,

Maryland Natural Resources Education, No. 28., 54 pp.

Association of National Estuary Program (ANEP) 2001. Indian River Lagoon Annual Report.

December, 112 pp.

Bahr, L.M. and W. P. Lanier 1981. The ecology of intertidal oyster reefs of the South Atlantic Coast:

a community profile. U.S. Fish Wildlife Services and Biological Service Program

FWS/OBS/-81/15:105.

Bartol, I.K., Mann, R., and M. Luckenbach 1999. Growth and mortality of oyster (Crassostrea virginica)

on constructed intertidal reefs. J. Exp. Mar. Bio. and Ecol. 237:157-184.

Bell, J.J. 2003. The importance of competitor identity, morphology, and ranking methodology to

outcomes in interference competition between sponges. Mar. Biol.143:415-426.

87

Boudreaux, M. L and L.J. Walters. Mytella charruana: a new invasive bivalve in the Indian River

Lagoon System, Florida. Submitted Nautilis.

Branch, G.M. 1984. Competition between marine organisms: ecological and evolutionary

implications. Ocean. Mar. Biol. Ann. Rev. 22:429-593.

Bros, W.E. 1987. Effects of removing or adding structure (barnacle shells) on recruitment to a

fouling community in Tampa Bay, Florida. J Exp. Mar. Biol. Ecol. 105:275-296.

Burrell V.G., Jr. 1982. Oyster Culture. in J.V. Hunter and E.E. Brown, eds. Crustacean and mollusk

aquaculture in the United States, AVI Publication Co., Westport, Conn, pp. 235-273.

Burrell, V.G. 1986. Species Profiles: Life History and Environmental Requirements of Coastal Fish

and Invertebrates- American Oyster. U.S. Fish and Wildlife Services Washington D.C.

Biological Rep. 82:11.57.

Bushek, D. 1998. Settlement as a major determinate of intertidal oyster and barnacle distributions

along a horizontal gradient. J .Exp. Mar. Biol. Ecol. 122:1-18.

Buss, L.W. 1981.Group Living, Competition, and the Evolution of Cooperation in a Sessile

Invertebrate Science 213:1012-1014.

88 Buss, L.W. and J.B.C. Jackson 1979. Competitive Networks: Nontransitive competitive relationships

in cryptic coral reef environments. Amer. Nat. 113:223-234.

Buss, L.W. 1981.Group Living, Competition, and the Evolution of Cooperation in a Sessile

Invertebrate Science 213:1012-1014.

Cake, E.W. Jr. 1983. Habitat Suitability Index models: Gulf of Mexico, American Oysters. U.S. Fish

Wild Serv. FWS/OBS-82/10/57.

Castagna, M. and P. Chanely 1973. Salinity tolerance of some marine bivalves from inshore and

estuarine environments in Virginia waters in the western mid-Atlantic coast. Melancholia

12:47-96.

Carriker, M.R. 1951. Ecological observations on the distribution of oyster larvae in New Jersey

estuaries. Ecol. Monog. 21:19-38.

Carriker, M.R. 1996. The shell and ligament. In the eastern oyster Crassostrea virginica eds. Kennedy ,

Newell and Eble, Maryland Sea Grant, College Park Maryland, pp. 75-168.

Chanely , P.E. 1957. Survival of some juvenile bivalves in water of low salinity. Proc. Nat. Shell.

Assosoation 48:52-65.

89 Coen, L.D., E. L. Wenner, D. M. Knott, B. W. Stender, N. H. Hadley, and M. Y. Bobo 1996.

Intertidal oyster reefs as critical estuarine environments: evaluating habitat use, development

and function. J. Shellfish Res. 15:490.

Coen, L., M. Luckenbach, and D.L. Breitburg 1999. The role of oyster reefs as essential fish habitat:

A review of current knowledge and some new perspectives. Amer. Fish. Society Symposium 22:

438-454.

Coen L and M.W. Luckenbach 2000. Developing success criteria and goals for evaluating

oyster reef restoration: ecological function or resource exploitation? Ecol. Eng. 15:323-343.

Connell, J.H. 1985. The initial settlement versus post-settlement in rocky intertidal communities. J.

Exp. Mar. Biol. Ecol. 93:11-46.

Connell, J.H. 1961. Effects of competition, predation of Thais lapillus, and other factors on natural

population the barnacle Balanus balaniodes. Ecol. Monog. 31:61-104.

Coon, S.L., Bonar, D.B. and R.M. Weiner 1985. Induction of settlement and metamorphosis of the

pacific oyster Crassostrea gigas (Thunberg) by L-dopa and catecholamines. J. Exp. Mar. Biol.

Ecol. 94: 211-21.

Coon, S .L., Walch, M., Fitt, M.K., Weiner, R.M. and D.B. Bonar 1990a. Ammonia induces

settlement behavior in oyster larvae. Biol. Bull. 179: 297-303.

90 Coon, S., Fitt, W.K. and D.B. Bonar 1990b. Competence and delay of metamorphosis in the pacific

oyster Crassostrea gigas. Mar.Biol. 106:379-87

Crabtree, R.E. and J.M. Dean. 1982. The structure of two South Carolina estuarine tide pool fish

assemblages. Estuaries 5:2-9.

Cranfield, H.J. 1973. Observation on the function of the glands on the foot on the pedivelagar

larvae of Orsrea edulis during settlement. Mar. Biol. 22:211-223.

Crisp D.J. 1976. Chemical factors inducing settlement in Crassostrea virginica (Gmelin). J. Anim. Ecol.

361:329-335.

Dalby, J. E. and C. M. Young 1993. Variable effects of ascidian competitors on oysters in a Florida

epifaunal community. J. Exp. Mar. Biol. Ecol. 167:47-57.

Dame, R.F. 1996. Ecology of marine bivalves an ecosystem approach. CRC Marine Science Series,

Boca Raton, Florida, USA., 256 pp.

Davis, H.C., and H. Hindu 1969. Effects of turbidity producing substances in sea water on eggs and

larvae of three genera of bivalve mollusks. Veliger 11:316-323

Davis, A.R. 1987. Variation in recruitment of the subtidal colonial ascidian Podoclavella cylindrical. J.

Exp. Mar. Biol Ecol. 106:57-71.

91 Eble, A.F. and R. Scro 1996. General Anatomy. In the Easter Oyster, Crassostrea virginica, edited by

Kennedy VS, Newell R, Eble AF, Maryland Sea Grant Program. College Park, MD, pp. 19-

74.

Fitt, W.K. and S.L. Coon. 1992. Evidence for ammonia as a natural cue for recruitment of oyster

larvae to oyster beds in a Georgia salt-marsh. Biol. Bull. 182:401-408.

Ford, S.E. and M.R. Tripp. 1996. Disease and Defense Mechanisms. In the eastern oyster

Crassostrea virginica, edited by Kennedy , REI, Newell and A.F. Eble, Maryland Sea Grant,

College Park Maryland, pp. 581-660.

Frechette, M and A.E. Aitken 1992. Interdependence of food and space limitation of a benthic

suspension feed: consequence for self-thinning relationships. Mar. Ecol. Prog. Ser. 83:55-62.

Frtiz, L.W., Lutz, R.A., Foote, M.A., VanDover, C.L. and J.W. Ewart 1984. Selective feeding and

grazing rates of oyster larvae on natural phytoplankton assemblages. Estuaries 7:513-518.

Gaines, S.D. , Brown, S. and J. Roughgarden. 1985. Spatial variations in larvae a concentration as a

case of spat variation in settlement. Ecology 67:267-272.

Galtsoff, P.S. 1964. The American Oyster Crassostrea virginica (Gmelin). Fish. Bull. 64, 484 pp.

Gardner, T. 1991. Mosquito Lagoon Aquatic Preserve Management Plan. Fl. Depart. Nat. Res..

183 pp.

92

Gosling, E. 2003. Bivalve molluscs: Biology, ecology and culture Oxford ; Malden, MA : Fishing

News Books, 453 pp.

Grizzle, R.E. 1990. Distribution and abundance of Crassostrea virginica (Gmelin,1791) (eastern oyster)

and Mercenaria spp. (quahogs) in a coastal lagoon. J. Shellfish Res. 9:347-358.

Grizzle, R.E., and M.W. Castagna. 1995. Final report – oyster reef monitoring program in

Canaveral National Seashore, National Park service, Titusville, FL. 14 pp.

Grizzle RE, Adams JR, and Walters LJ 2002. Historical changes in intertidal oyster (Crassostrea

virginica) reefs in a Florida lagoon potentially related to boating activities. J. Shellfish Res. 2:

749-756.

Grosberg, R. K. 1981. Sperm-mediated gene flow and the genetic structure of a population of the colonical ascidian Botryllus schlosseri. Evolution 45:130-142.

Gunter G 1979. Studies of the southern oyster borer, Thais haemastoma. Gulf Research

Reports 6: 249-260.

Gurevitch, J., J.A. Morrison and L.V. Hedges 2000. The interaction between competition on and

predation: a metanaylsis of field experiments. American Naturalist 155:435-453.

93 Hadfield, M.G. 1978. Metamorphosis in marine molluscan larvae: an analysis of stimulus and

response. In: Settlement and Metamorphosis of Marine Invertebrate Larvae. New york

Press, pp 165-175.

Hart, A.W. 1994. A Boater’s Guide to the Indian River Lagoon. 85 pp.

Haven, D.S. and R. Morlaes-Alamo. 1966. Filtration of particles from suspension by the American

oyster, Crassostrea virginica . Biol. Bull. 139:248-264.

Indian River Lagoon Comprehensive Conservation and Management Plan 1996. Indian

River Lagoon National Estuary Program. Nov. Report, 193 pp.

Ingle, R.M.F and C.E. Dawson, Jr. 1950. Variation in salinity and its relation to the Florida oyster.

Pro. Nat. Shell. Ass. 23:16-19.

Jackson, J. B. C. 1979. Overgrowth competition between encrusting cheilostome ectoprocts in a

Jamaican cryptic reef environment. J. Anim. Ecol., 48:805-823.

Jackson, J.C., M. X. Kirby, W. H. Berger, K. A. Bjorndal, L. W. Botsford,B. J. Bourque,R. H.

Bradbury, R. Cooke, J. Erlandson, J. A. Estes, T. P. Hughes, S. Kidwell, C. B. Lange, H. S.

Lenihan, J. M. PandolÞ, C. H. Peterson, R. S. Steneck, M. J. Tegner, and R. R. Warner. 2001.

Historical overfishing and the recent collapse of coastal ecosystems. Science 293:629-639.

94 Kennedy, V. S. and L. L. Breisch 1980. Maryland's Oysters Research and Management Maryland Sea

Grant, University of Maryland College Park, Maryland, 286 pp.

Kennedy, V.S., Newell RIE and A.F. Eble 1996. The Eastern Oyster. Maryland Sea Grant,

College Park, MD, 772 pp.

Kennedy, V.S., and L.P andford 1999. The morphology and physical oceanography of unexploited

oyster reefs in North America. Oyster Reef Habitat Restoration: A Synopsis and Synthesis of

Approaches, VIMS Press, Gloucester Point, VA.

Keough, M.J. and B.J. Downs 1982. Recruitment of marine invertebrates: the role of active larval

choice and early morality. Oecoligica 54:348-352.

Korringa, P. 1952. Recent Advances in Oyster Biology. The Quarterly Review of Biology 27:266-308.

Lenihan, H. S. 1999. Physical-biological coupling on oyster reefs: how habitat structure

influences individual performance. Ecol. Monog. 69:251-276

Lenihan H.S. and F. Micheli. 2001. Soft-sediment communities. In Marine Community Ecology,

edited by M.D. Bertness S.D. Gaines, and M.E. Hay, Sinaeur Associates, Inc., Sunderland,

M.A., pp 253-287.

95 Lewis, CA. 1978. A review of substratum selection in free-living and symbiotic Cirripeds. In

Settlement and Metamorphosis of Marine Invertebrate Larvae. New York: Elsevier, 1978.

pp 147-159.

Levitan, D.R. 1995. The ecology of fertilization in free-spawning invertebrates. In Ecology of

Marine Invertebrate Larvae, edited by L. McEdward, CRC press, Boca Raton FL., pp.123-

156.

Lohse, D. 2002. Relative strengths of competition for space and food in a sessile filter feeder. Biol.

Bull. 203:173-180.

Loosanoff, V.L., and C.A. Nomejko 1946. Feeding of oysters in relation to tidal stages and to

periods of light and darkness. Biol. Bull. 90:244-264.

Loosanoff, V.L. and F.D. Tommers 1948. Effect of suspended silt and other substances in the gut

of feeding in oysters. Science 107:69-70.

Loosanoff, V.L. 1953. Behavior of oysters in water of low saltines. Proc. Nat. Shell. Ass. 43:135-51.

Loosanoff, V.L. 1962. Effects of turbidity on some larval and adult bivalves. Proc. Gulf Carib. Fish.

14:80-93.

Loosanoff, V. L. 1965. Gonad development and discharge of spawn in oysters of

Long Island Sound. Biol. Bull. 129: 546-561.

96

MacKenzie, C.L. 1979. Sea anemone predation on larval oysters in Chesapeake Bay

(Maryland). Proc. Nat. Shellfish Ass. 67:113-117.

MacKenzie, C.L. 1983. To increase oyster production in the northeastern United States.

Marine Fish. Rev. 45:1-22.

MacKenzie, C.L. 1996a. History of oystering in the United States and Canada, featuring the eight

greatest oyster estuaries. Marine Fish Review. 58:1-87.

MacKenzie, C.L. 1996b. Management on natural populations. The Eastern Oyster,

Crassostrea virginica. Edited by VS Kennedy, RIE Newell, and AF Ebele. Maryland

Sea Grant, College Park, MD, pp 707-719.

Mackie, G. 1984. Bivalves. In The Mollusca Vol 7., edited by AS Tompa, NH Verdonk and J.A.M.

Van den Biggelaar, Academic Press, New York, pp. 351-418.

Mangum, D. C., B. P. Shepard, and J. C. Williams 1972. Methods of controlling marine fouling in

desalination plants. Proceedings: Third International Congress on Marine Corrosion and

Fouling pp 357-364.

97 Menzel, R.W. 1991. Cultural practices and production of the American oyster with emphasis on the

south Atlantic and Gulf of Mexico. In Estuarine and Marine Bivalve Mollusk Culture, edited

by W. Menzel, CRC Press, Boston, pp 25-34.

Micheli, F. and C.H. Peterson 1999. Estuarine vegetative habitats as corridors for predator

movement. Conservation Biology 13:869-881.

Mook, D.H.. 1976. Studies on fouling invertebrates in the Indian River. I . Seasonality of settlement.

Bulletin Mar. Sci. 26: 610-615.

Mook, D.H. 1977. Studies on fouling organisms in the Indian River Lagoon 2. Effect of Modulus

modulus. Nautilus 91:134-136.

Mook, D. 1980. Seasonal Variation in species composition of recently settled fouling communities

along an environmental gradient in the Indian River Lagoon, Florida.

Estuarine and Coastal Science 2: 573-581.

Mook, D.H. 1981. Effects of disturbance and initial settlement on fouling community structure.

Ecology 62:522-526.

Mook, D. 1983. Indian River Lagoon fouling organisms, a review. Florida Scientist 46:162-167.

98 Nelson, K.A., Leonard,L,A., Martin H.P., Alphin T.D., and M.A. Mallin 2004. Using transplanted

oyster (Crassostrea virginica) beds to improve water quality in small tidal creeks: a pilot study.

J. Exp. Mar. Biol Ecol. 298:347-368.

Newell R.L. 1988. Ecological changes in Chesapeake Bay: are they the result of overharvesting the

American oyster? Chesapeake Bay Research Consortium, Baltimore, MD. 129:536-546.

Nowell A.R.M. and P.A. Jumars 1984. Flow environments of aquatic benthos. Ann. Rev. Ecol. Syst.

15: 303-328.

Okamura, B. 1986. Group living and the effects of spatial position in aggregation of Mytilus edulis.

Oecologia 69: 341-347.

Osman, R.W., R.B. Whitlach, and R.N. Zajac 1989. Effects of resident on recruitment into a

community: larva settlement versus post settlement mortality in the oyster Crassostrea virginica

Mar. Ecol. Prog. Ser. 54:61-73.

Osman, R.W., R.B. Whitlach , R.J. Malatesta and R.N. Zajac 1990. Ontongenic changes in tropic

relationship and their effects on recruitment. In: Trophic Relationships in the Marine

Environment, Aberdeen Press University Press, pp 117-129.

Paine R.T. 1969. Pisaster-Tegula interaction – prey patches, predator food preference, and intertidal

community structures. Ecology 50:950.

99 Peterson C.H. and R. Black 1987. Resource depletion by active suspension feeders on tidal mudflats:

influence of local density and tidal elevation. Limno. Ocean. 32:143-166.

Provancha, J.A., C.R. Hall, and D.M. Oddy. 1992. Mosquito Lagoon environmental resources

inventory. NASA Technical Memorandum 107548. The Bionetics Corporation. Kennedy

Space Center, Florida.

Quale O.B. 1969. Pacific oyster culture in British Columbia .Fish. Res. Canada Bull. 169. 193 pp.

Ritchis T.P and R. W. Menzel 1969. Influences of light on larval settlement of the American oysters.

Pro. Nat. Shell. Ass. 59:116-120.

Roegner, C.G. 1991. Temporal analysis of the relationship between settlers and early recruits of the

oyster Crassostrea virginica (Gmelin). J. Exp. Mar. Biol Ecol. 151:57-69.

Roughgarden J, Gaines, S., and H. Possingham. 1988. Recruitment dynamics in complex life cycles.

Science 241:1460-1466.

Rumrill, S.S.1990. Natural morality of marine invertebrate larvae. Ophelia 32:163-198.

Ryland, J.S., 1977. Taxes and tropisms of Bryozoans. In Biology of Bryozoans, edited by R.M.

Woollacott & R.L. Zimmer, pp. 411-436.

100 Scheltema, R.S., 1974. Biological interactions determining larval settlement of marine invertebrates.

Thalassia Jugoslavica, 10:263-296.

Seliger HH, Boggs JA, Rivkin RB, and W. H. Biggley 1982. The transport of oyster larvae in an

estuary. Mar. Biol.. 17:57-72.

Sellers, M.A. and J.G. Stanley 1984. Species profiles: Life history and environmental requirements of

coastal fishes and invertebrates (North Atlantic)- American oyster. U.S. Fisheries and

Wildlife Services OBS-83/11.23 U.S. Army Corps of Engineers, TR EL-82-4. 15 pp.

Shelbourne, J.E. 1957. The 1951 oyster stock in the River Crouch and Roach, Essex. Fish. Invest.,

London, 21:1-25.

Smithsonian Institution 2001. Indian River Lagoon Species Inventory Report. Website:

http://www.sms.si.edu/irlspec/index.htm.

Sotka, E.E. 2005. Local adaptation in host use among marine invertebrates. Ecology Letters 8:448-459.

Surge, D.M. , Lohmann, K.C. and G.A. Goodfriend 2003. Reconstructing estuarine conditions:

oyster shells as records of environmental change, Southwest Florida. Estuarine Coastal and

Shelf Sciences 57:737-756.

101 Tamburri MN, Zimmer-Faust RK, and M.L. Tamplin 1992. Natural sources and properties of

chemical inducers mediating settlement of oyster larvae: A re-examination. Biol. Bull. 83:327-

338.

Tamburri MN, Finelli CM, Wethy DS and R.K. Zimmer-Faust 1996. Chemical induction of larval

settlement behavior in flow. Biol. Bull.. 191:367-373.

Tolley, S.G., A. K. Volety, and M. Savarese 2005. Influence of salinity on the habitat use of oyster

reefs in three southwest Florida estuaries. J. Shellfish Res. 24:127-138.

Turner EJ, Zimmer-Faust RK, Palmer MA, Luckenbach M, and N.D. Pentchef 1994. Settlement of

oyster (Crassostrea virginica) larvae: Effects of water flow and a water-soluble chemical cue.

Limno. Oceano. 39:1579-1593.

Underwood A.J. and P.G. Fairweather 1989. Supply side ecology and benthic marine assemblages.

Trends in Ecol. Evo. 4:16-20.

Underwood,A.J. and M.J. Keough 2001. Supply-side ecology. The nature and consequences in

variations in recruitment of intertidal organisms. In Marine Community Ecology, Sinaure

Ass., 560 pp.

Wall, L.M. 2004. Recruitment and restoration for the oyster Crassostrea virginica on intertidal reefs in

areas with intense boating activity.(Master's thesis, University of Central Florida, 2004).

Masters Abstracts International.

102

Wall, L.M. , L. J. Walters, R.E. Grizzle, and P.E. Sacks. In press. Recreational boating activity and its

impacts on the recruitment and survival of the oyster Crassostrea virginica on intertidal reefs in

Mosquito Lagoon, Florida. J. of Shellfish Research.

Wallace, R.K. 2001. Cultivating Crassostrea virginica. Southern Region Aquaculture Center Publication

No. 432. 4 pp.

Walters LJ 1992. Field settlement locations on subtidal marine hard substrata: Is active larval

exploration involved? Limno. Oceano. 37:1101-1107.

Walters LJ, Roman A, Stiner J, and Weeks D 2001. Water Resource Management Plan, Canaveral

National Seashore. Titusville, FL: National Park Service, Canaveral National Seashore. 224

pp.

Wells, H.W. 1961. The fauna of oyster reefs with special reference to the salinity factor. Ecol. Mono.

60:449-469.

Wenner, E., H.R. Beatty and L. Coen 1996. A quantitative system for sampling nekton on intertidal

oyster reefs. J. Shellfish Res. 15:769-775.

White, M.E. and E.A.Wilson 1996. Predators, pests and competitors. In The Eastern Oyster

Crassostrea virginica , edited by Kennedy R.E.I, Newell and A.F. Eble Maryland Sea Grant,

College Park Maryland, pp 559-579.

103 Young, C.M. 1990. Larval predation by epifauna on temperate reefs: scale, power, and the scarcity of

measurable effects. Aust. J. Ecol. 15:413-426.

Zajac, R.N., R.B. Whitlach and R.W. Osman 1989. Effects of inter-specific density and food supply

on survivorship and growth of newly settled benthos. Mar. Ecol. Prog. Ser. 56:127-132.

Zimmer-Faust, R.K. and M.N. Tamburri 1994. Chemical identity and ecological implications of a

waterborne, larval settlement cue. Limn. Ocean. 39:1075-1087.

Zullo, V. A.. 1963. A preliminary report on systematics and distribution of barnacles (Cirripeda) of

Cape Cod region. Systemic-Ecology Program, Mar. Bio Laboratory, Woods Hole Mass., 33

pp.

104