Open Collections

UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Invertebrate fouling community composition associated with Pacific oyster (Crassostrea gigas) suspended… Switzer, Soleil Elana 2010

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Item Metadata

Download

Media
24-ubc_2010_spring_switzer_soleil.pdf [ 1.49MB ]
Metadata
JSON: 24-1.0069194.json
JSON-LD: 24-1.0069194-ld.json
RDF/XML (Pretty): 24-1.0069194-rdf.xml
RDF/JSON: 24-1.0069194-rdf.json
Turtle: 24-1.0069194-turtle.txt
N-Triples: 24-1.0069194-rdf-ntriples.txt
Original Record: 24-1.0069194-source.json
Full Text
24-1.0069194-fulltext.txt
Citation
24-1.0069194.ris

Full Text

Invertebrate fouling community composition associated with Pacific oyster (Crassostrea gigas) suspended tray culture  by  SOLEIL ELANA SWITZER B.Sc., Malaspina University-College, 2004  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE  in  THE FACULTY OF GRADUATE STUDIES  (Animal Science)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  March 2010  © Soleil Elana Switzer, 2010  ABSTRACT Fouling organisms associated with suspended oyster aquaculture can significantly increase operational costs for growers and significantly decrease product marketability. Currently, there is little information available on the fouling communities present on deep-water, suspended tray Pacific oysters (Crassostrea gigas). In general, the industry practice in tray production of Pacific oyster is to use plastic trays, but a new polyvinyl-coated metal tray, which accommodates higher oyster densities, has recently become available. The objectives of this thesis were firstly, to assess whether differences exist in the extent of tray fouling between the 2 tray types using abundance, species richness, biomass, and dominance (univariate analyses) and the Bray-Curtis Dissimilarity Coefficient (multivariate, cluster analysis) as measures (Chapter 2) and secondly, to describe variation in the fouling organisms present on the oysters in the 2 tray types using the same univariate and multivariate analyses (Chapter 3). This study took place on a commercial oyster farm between the months of October 2006 and October 2007. Overall, there was little difference in the extent of fouling communities between the tray types, although a few species were present on plastic trays only, but in very low abundance. The fouling communities associated with the trays and oysters were studied in the winter, spring, summer (tray only) and fall to determine seasonal variation in the fouling communities. Abundance, species richness, biomass and dominance in the tray fouling communities were affected by season with the greatest values in July and the lowest values in January. Season also influenced the oyster fouling communities. Abundance, biomass and dominance of the fouling communities on the oysters was higher in October, while species richness was lowest in January compared to April or October. The seasonal changes observed in the fouling communities on both the trays and oysters were driven largely by a few species. High abundances of Caprella mutica and Mytilus sp. on the trays in July could impact oyster growth through the reduction of water flow through the trays or, in the case of Mytilus sp., through direct competition with the oysters for food and/or space.  ii  TABLE OF CONTENTS  ABSTRACT ..................................................................................................................................... ii TABLE OF CONTENTS .............................................................................................................. iii LIST OF TABLES .......................................................................................................................... v LIST OF FIGURES ....................................................................................................................... vi ACKNOWLEDGEMENTS ........................................................................................................ viii CO-AUTHORSHIP STATEMENT ............................................................................................. ix  CHAPTER ONE. Introduction 1.1 RESEARCH TOPIC........................................................................................................... 1 1.1.1 Suspended oyster culture operations as substrate for fouling communities ......... 1 1.1.2 Factors affecting fouling community composition ................................................... 3 1.1.3 Treating fouling associated with bivalve aquaculture ............................................. 7 1.2 THESIS OVERVIEW ........................................................................................................ 8 LITERATURE CITED .................................................................................................................. 9  CHAPTER TWO. Effects of season and tray type on fouling communities associated with oyster trays 2.1 INTRODUCTION ............................................................................................................ 17 2.2 METHODS ........................................................................................................................ 18 2.2.1 Study site and experimental design ......................................................................... 18 2.2.2 Field methods ............................................................................................................ 20 2.2.3 Laboratory methods ................................................................................................. 21 2.2.4 Data analysis .............................................................................................................. 22 2.3 RESULTS .......................................................................................................................... 24 2.3.1 Composition of fouling communities on oyster-culture trays ............................... 24 2.3.1.1 Abundance .............................................................................................................. 24 2.3.1.2 Biomass ................................................................................................................... 27 2.3.1.3 Species richness ...................................................................................................... 28 2.3.1.4 Swartz Dominance Index (SDI) .............................................................................. 30 2.3.1.5 Community analyses ............................................................................................... 31 2.3.2 Seasonal and spatial variation in water column parameters ................................ 33 2.4 DISCUSSION .................................................................................................................... 35 2.5 CONCLUSIONS ............................................................................................................... 40 LITERATURE CITED ................................................................................................................ 42  iii  CHAPTER THREE. Seasonal comparison of invertebrate fouling communities associated with Crassostrea gigas cultured in two different tray types  3.1 INTRODUCTION ............................................................................................................ 47 3.2 METHODS ........................................................................................................................ 48 3.2.1 Study site and experimental design ......................................................................... 48 3.2.2 Field methods ............................................................................................................ 49 3.2.3 Laboratory methods ................................................................................................. 50 3.2.4 Data analysis .............................................................................................................. 51 3.3 RESULTS .......................................................................................................................... 52 3.3.1 Composition of fouling communities on tray cultured oysters ............................. 52 3.3.1.1 Abundance .............................................................................................................. 53 3.3.1.2 Biomass ................................................................................................................... 55 3.3.1.3 Species richness ...................................................................................................... 57 3.3.1.4 Swartz Dominance Index (SDI) .............................................................................. 58 3.3.1.5 Community analyses ............................................................................................... 59 3.4 DISCUSSION .................................................................................................................... 61 3.5 CONCLUSIONS ............................................................................................................... 66 LITERATURE CITED ................................................................................................................ 68  CHAPTER FOUR. Summary and conclusions 4.1 Seasonal variation in fouling community composition .................................................. 72 4.2 Tray-type and fouling community composition ............................................................. 73 4.3 Research implications ....................................................................................................... 73 4.4 Future research ................................................................................................................. 74 LITERATURE CITED ................................................................................................................ 76  iv  LIST OF TABLES  Table 2.1. Comparison of key differences between the plastic (PL) and polyvinyl (PV) experimental trays…………………………………………………………………………..20 Table 2.2. Number of replicate samples for each fouling community (date and tray types).........23 Table 2.3. Summary of two-factor ANOVA results (p-values) for the 22 major taxonomic groups for abundance (log (x+0.1) transformed number individuals per tray), biomass (log (x+0.1) grams per tray) transformed and, species richness (number species per tray), for two factors (date and tray type) and the interaction between the two factors...........................................25 Table 2.4. Summary of two-factor ANOVA results for total abundance (log (x+0.1) transformed total number of individuals per tray), biomass (log (x+0.1) transformed total biomass per tray) and species richness (total number of species per tray) for 2 factors (date and tray type) and for the interaction between the 2 factors……………………………………………….25 Table 3.1. Summary of two-factor ANOVA results (p-values) for the 21 major groups for abundance (log (x+0.1) transformed) (number individuals per m2), biomass (log (x+0.1) transformed) (g per m2) and species richness (number species per m2) for 2 factors (date and tray type) and the interaction between the 2 factors………………………………………..54 Table 3.2. Summary of two-factor ANOVA results for abundance (log (x+0.1) transformed) (total number individuals per m2), biomass (log (x+0.1) transformed) (total g per m2) and species richness (total number species per m2) for 2 factors (date and tray type) and for the interaction between the 2 factors………………………………………………………...….54  v  LIST OF FIGURES  Figure 2.1. Location of oyster farm, experimental raft and water parameter stations. (a) Oyster farm, (b) Oyster farm in the embayment, the experimental raft and the stations used in the collection of water column parameter data…………………………………………………19 Figure 2.2. Experimental oyster culture trays. (a) “traditional” plastic oyster trays stacked seven trays high (note gaps between trays), (b) polyvinyl oyster trays stacked ten trays high (notice trays fit tightly together), (c) plastic oyster tray (PL) and (d) new metal oyster tray with polyvinyl coating (PV)………………………………………………………………...19 Figure 2.3. Seasonal changes in fouling communities on PL and PV trays. (a) Mean total abundance (log transformed) per fouling community (number of individuals per tray + SE), (b) Mean total biomass (log transformed) per fouling community (g per tray + SE) and (c) Mean total species richness per fouling community (number of species per tray + SE)…...26 Figure 2.4. Seasonal mean abundance (log transformed) per fouling community (number individuals per tray) on suspended PL and PV trays for (a) Class Bivalvia, (b) Class Demospongiae and (c) Class Polychaeta…………………………………………………...26 Figure 2.5. Seasonal mean biomass (log transformation) per tray fouling community (g per tray + SE) on suspended PL and PV trays for (a) Class Bivalvia, (b) Class Malacostraca and (c) Class Asteroidea…………………………………………………………………………….28 Figure 2.6. Seasonal mean species richness per fouling community (number species per tray) on suspended PL and PV trays for (a) Class Malacostraca, (b) Class Polychaeta and (c) Class Gastropoda………………………………………………………………………………….29 Figure 2.7. Mean dominance per fouling community on suspended PL and PV trays (SDI per tray + SE)…………………………………………………………………………………...30 Figure 2.8. Hierarchical dendrogram showing the relative similarities of fouling communities using the Bray-Curtis dissimilarity index values of species abundance. Each letter (A through G) is associated with a cluster linkage……………………………………………..32 Figure 2.9. Hierarchical dendrogram showing the relative similarities of fouling communities using the Bray-Curtis dissimilarity index based on species biomass. Each letter (A through G) is associated with a cluster linkage……………………………………………………...32 Figure 2.10. Water column profiles in January, April, July and October at the raft centre (C) station. (a) Temperature, (b) pH, (c) salinity and (d) dissolved oxygen profiles were created using Sonde data, (e) chlorophyll and (f) turbidity (TSS) profiles were created using labanalyzed Niskin water samples. Chlorophyll values for January are hidden by the October symbols……………………………………………………………………………………..34  vi  Figure 3.1. Mean total oyster surface area per fouling community on oysters suspended in PL and PV-trays (m2 + SE)…………………………………………………………………….53 Figure 3.2. Seasonal changes in fouling communities on oysters suspended in PL and PV-trays. (a) Mean total abundance (log transformed) per fouling community (number of individuals per m2 + SE), (b) Mean total biomass (log transformed) per fouling community (g per m2 + SE) and (c) Mean total species richness per fouling community (number of species per m2 + SE)…………………………………………………………………………………………..55 Figure 3.3. Seasonal mean abundance (log transformed) per fouling community (number individuals per m2) on oysters suspended in PL and PV-trays for (a) Class Polychaeta, (b) Class Malacostraca and (c) Class Demospongiae…………………………………………..55 Figure 3.4. Seasonal mean biomass (log transformation) per fouling community (g per m2) on oysters suspended in PL and PV-trays for (a) Class Demospongiae, (b) Class Gymnolaemata and (c) Class Holothuridea………………………………………………...56 Figure 3.5. Seasonal mean species richness per fouling community (number species per m2) on oysters suspended in PL and PV-trays for (a) Class Malacostraca, (b) Class Polychaeta and (c) Class Gymnolaemata……………………………………………………………………57 Figure 3.6. Seasonal mean dominance per fouling community on oysters suspended in PL and PV-trays (SDI per community + SE)……………………………………………………….59 Figure 3.7. Hierarchical cluster dendrogram showing the relative dissimilarities of fouling communities using an unweighted pair-group mean average sort algorithm based on BrayCurtis dissimilarity index values of species abundance data. Each letter (A through G) is associated with a cluster linkage……………………………………………………………60 Figure 3.8. Hierarchical cluster dendrogram showing the relative dissimilarities of fouling communities using the Bray-Curtis dissimilarity index based on species biomass data. Each letter (A through G) is associated with a cluster linkage…………………………………...60  vii  ACKNOWLEDGEMENTS The major financial support for this study was provided by the Natural Sciences and Engineering Research Council of Canada (NSERC) industrial post-graduate scholarship program made possible through the generous contributions by the Evening Cove Oysters (ECO) of Nanaimo. ECO was a tremendous help in making this project happen by providing a field site, transportation to and from the site, the equipment and oysters to set-up the field experiment and most importantly continuous support in ensuring the study was successful. I would also like to thank the Centre for Shellfish Research (CSR) at Vancouver Island University (VIU) for their additional financial support and in-kind contribution of field and lab equipment. I express appreciation to my committee members Drs. Scott McKinley (supervisor), Brenda Burd and Nina von Keyserlingk for guidance and support. A special thanks to my co-supervisor Dr. Penny Barnes who provided financial, academic and moral support through this very ambitious project. The staff at the CSR, – in particular Don Tillapaugh – VIUs’ Fisheries and Aquaculture Department and VIUs’ Biology Department were extremely supportive in providing students and staff to assist me in the field. A special thanks to all my field helpers –Kate, Simon, Edith, Johanne, Spencer, Christian, Grant, Cam, Mike, Jackson, Julie, Graham, Aaron, Jacqueline, Erin, Desiree, Larissa and Nicky - for long days in the sun, rain and snow to perform the tedious job of removing fouling from oysters, without all of you I could have never done it. Thanks to Nadia Plamondon, Kate Rolheiser and Sandy Lipovsky for hanging in there, I know spending countless hours sorting fouling invertebrates is not everyone’s idea of fun. A special thanks to my close friends and family who were forced to help when there was no one else, who listened to problems and pains, who accepted that I could not be a part of most things because I had to work and most of all held me up and pushed me along when I thought I could not go any further.  viii  CO-AUTHORSHIP STATEMENT I designed this research project, conducted the research, analyzed the data, and prepared the manuscript. As co-authors on the two manuscripts, Penny Barnes and Robert McKinley provided direction in the experimental design, advice on data analyses, and aided in the preparation of the manuscripts.  ix  CHAPTER ONE. Introduction 1.1  RESEARCH TOPIC  In the marine environment, the submersion of an artificial substrate into the water column promotes colonization of marine life, including bacteria and invertebrates, in a process termed fouling. Fouling has become particularly important due to its impact in 2 main sectors: marine transport and marine aquaculture (Wahl, 1989). Fouling on the hulls of ships increases drag and decreases speed. In addition, fouling organisms can be transported large distances, otherwise unattainable through natural recruitment processes, resulting in the potentially problematic introduction of invasive species (Aleem, 1957; Railkin, 2004). In marine aquaculture, particularly suspended bivalve aquaculture, diverse invertebrate fouling communities form through colonization of both abiotic (farming equipment) and biotic (cultured bivalves) substrates. Due to the abundance of available substrate in a commercial-size operation, excessive fouling can impact the growth and survival of the cultured bivalves, particularly juveniles, directly through competition (space and food) and predation (Arakawa, 1990; Anderson, 1999; Miron et al., 2002; LeBlanc et al., 2003), as well as indirectly by restricting water circulation in nets or trays or by extra handling of oysters when removing or treating fouling (Parsons and Dadswell, 1992; Claereboudt et al., 1994; Taylor et al., 1997). In addition, fouling on aquaculture equipment and cultured bivalves increases the weight and the drag of the culture system, which in turn affects the stability and buoyancy of the operation (Enright, 1993; Erbland and Ozbay, 2008; Rodriguez and Ibarra-Obando, 2008).  1.1.1 Suspended oyster culture operations as substrate for fouling communities While Canadian bivalve aquaculture may make only a small contribution to the global market, Canada’s industry has huge potential for expansion. Currently, Canadian shellfish growers culture 4 main bivalves (mussels, oysters, clams and scallops) and the culture of the Pacific oyster (Crassostrea gigas) in British Columbia (BC) contributes about 17% of Canada’s total gross bivalve tonnage (DFO, 2008). The Pacific oyster is a non-native species brought to Canada specifically for aquaculture purposes because the slow growth of the native Olympia oyster (Ostrea conchapila) makes it unsuitable for culture (BCSGA, 2009). Oyster aquaculture in BC is dynamic, with culture methods continually evolving to address changes in the market and sitespecific characteristics, to ensure that growers maximize cost benefits. BC oyster growers utilize different culture methods based on the hydrographic attributes of their intertidal or deep-water  1  lease, the intended market (shucked or half-shell) and their operation capability (BCSGA, 2009). Depending on the intended market, oysters must meet particular standards. For example, oysters that are grown to be shucked can generally vary in shape and size and the shells can have some fouling but oysters grown for the half-shell market must meet a much higher appearance standard. Most oysters intended for the half-shell market will grow-out in deep-water tray systems, known to produce fast growing, high quality oysters. Deep-water tray systems use rafts or long-lines anchored in the subtidal zone to support stacks of trays suspended in the water column. Trays must be stacked and strapped together in a way that ensures they remain vertical in the water column (BCSGA, 2009). Oyster culture trays have been modified over the years to allow adequate flow and space to each oyster and more recently, to reduce fouling. Two tray types are commonly used in BC for growing oysters, the “traditional” plastic tray and a newer metal tray with polyvinyl coating. Aquaculture tray systems are labour intensive because oysters must be graded (densities reduced) as they grow to guarantee maximum production. When trays are lifted to reduce oyster densities, most farms will tumble oysters and either transfer oysters to clean trays or remove the major fouling from the trays (BCSGA, 2009). In the tumbling process, oysters are placed in a rotating inclined cylinder where they are thrown around while moving from one end of the cylinder to the other. The tumbling process removes excess shell (frills) and some fouling organisms. The removal of excess shell allows the oyster to concentrate energy stores to meat growth instead of shell growth, thus creating the perfect cupped oyster. The deep-water, tray culture of oysters provides an ideal environment for the settlement and growth of diverse fouling communities because the numerous oyster trays, and the oysters themselves, create a complex three-dimensional structure, similar to a reef, with abundant substrate suitable for invertebrate settlement and attachment (DFO, 2006). Oyster rafts create a baffle effect whereby the suspended stacks collectively dissipate wave energy, modifying hydrodynamics and creating a calm environment within the raft that is ideal for larval settlement and that traps plankton for filter-feeders (Wahl, 1989; Bourget et al., 1994; Guiterrez et al., 2003). These rafts also provide an abundance of attached prey species for predators and the complexity of oyster rafts, particularly the vertical stacks, can provide protection from visual predators (McKindsey et al., 2006). The impact fouling communities have on cultured bivalves can be minimized with labour-intensive regular cleaning of the gear and the bivalves.  2  1.1.2  Factors affecting fouling community composition  The composition of fouling communities at bivalve aquaculture sites depends on the biotic and abiotic substrate available for colonization (species cultured, culture methods), the location of the site (spatial variation in water parameters, currents, larval recruitment) and the time of year and length of time the substrate is deployed (available colonists). In the marine environment, the availability of natural, hard substrate for settlement by sessile marine organisms is a limiting factor (Lesser et al., 1992). Naturally-occurring hard substrates include rock and mollusc shells. Bivalve aquaculture operations increase the availability of biotic substrate, by introducing large densities of shells for colonization (both temporarily (cultured bivalve shells) and permanently (lost or discarded bivalve shells)). In addition to biotic substrate, the introduction of abiotic/artificial semi-permanent (docks, aquaculture rafts) or temporary (boats, aquaculture nets or cages) substrates into intertidal and subtidal coastal areas substantially increases substrate availability. Most marine fouling organisms are sessile invertebrates and therefore fouling community composition is largely dependent on the colonization of available substrates. Fouling communities become established on a new substrate through a series of overlapping phases during the colonization process. The phases of marine colonization, more often termed succession in community ecology, are biochemical conditioning, bacterial colonization, unicellular fouling and multicellular fouling (Aleem, 1957; Wahl, 1989; Railkin, 2004). Biochemical conditioning begins immediately after a substrate enters seawater and involves the adsorption of dissolved chemicals from the seawater to the substrate. Within an hour of submersion, bacteria colonize the substrate and form a film. Unicellular species, such as protozoans and diatoms begin settling on the film within days. The last stage of colonization, multicellular fouling, may begin as early as several days after submersion of the substrate, initiated by larvae and algal settlement (Wahl, 1989). Multicellular, particularly invertebrate, presence in a fouling community is largely dependent upon the recruitment (settlement and metamorphose) of planktonic larvae from the water column on to a suitable substrate (Osman, 1977; Bonar et al., 1986). Invertebrate larvae have developed unique species-specific selection methods for determining substratum suitability. Most larvae have the ability to detect specific cues from the presence or absence of particular biochemical compounds or species (including conspecifics) occupying the substratum, as well as cues from the environment (Bayne, 1965; Osman, 1977; Bonar et al., 1986; Walters and Wethey, 1991; Woodin, 1991; Qian, 1999; Bullard et al., 2004).  3  It has been documented that some larvae have a preference for substrate of a particular colour or texture (Dahlem et al., 1984; Walters, 1992; Callow and Fletcher, 1994). For example, Anderson and Underwood, (1994) demonstrated that invertebrate species colonize hard, porous materials (e.g. concrete, plywood) in greater numbers than they do hard smooth, substrates (e.g. fibreglass, aluminum). In addition, it has been suggested that smooth artificial surfaces also support greater abundances of some non-native ascidians than do porous surfaces (Tyrell and Byers, 2007). Substrate rejection is risky to most species, especially non-feeding larvae, because the proximity of an acceptable substratum is unknown. Movement to a new substrate can occur by rolling, crawling or swimming small distances, or by returning to the water column to travel longer distances (Woodin, 1991). The colonization and composition of marine fouling communities have been studied using an array of artificial substrates such as glass, plexiglass, asbestos, cement, asbestos-cement, plastic, wood, ceramic/clay, marble/slate/granite, metal, vinyl acetate and polyvinyl chloride (for example Aleem, 1955; Dean and Hurd, 1980; Chalmer, 1982; Okamura, 1986; Anderson and Underwood, 1994; Brown and Swearingen, 1998; Tyrell and Byers, 2007). Very few of these studies examined the fouling community composition in detail, focusing rather on the presence of dominant species and their role in community succession, the role of particular species as competitors in fouling communities, or the effects of disturbance patterns on community assemblages. Fouling communities have been documented in the setting of bivalve aquaculture for the following bivalves: the great Atlantic scallop Pecten maximus (Ross et al., 2004), the giant scallop Placopecten magellanicus (Claereboudt et al., 1994), the tropical scallop Euvola ziczac (Lodeiros and Himmelman, 2000), the blue mussel Mytilus edulis (Lesser et al., 1992; LeBlanc et al., 2003), the pearl oysters Pinctada fucata (Murad and Mohammad, 1976; Guenther et al., 2006), Pinctada imbricata (Lodeiros and Garcia, 2004), and Pinctada maxima (Taylor et al., 1997), the Pacific oyster C. gigas (Arakawa, 1990; Mazouni et al., 2001; Rodriguez and Ibarra-Obando, 2008) and the Eastern oyster Crassostrea virginica (Erbland and Ozbay, 2008). Comparison of the results of these studies is extremely difficult not only because of the variety of substrates used, but also because of the variation in location and climate, season, methodology and the geographic variation in larval recruitment (temporal and spatial). In addition, the variability in the composition and size of the biotic substrate material (shells) and the variation in morphology and  4  life history of the substrate bivalves means comparison of these studies is severely limited (Underwood and Chapman, 2006). Besides larval recruitment and substrate type, other factors influencing the development of a fouling community include: disturbances (e.g. competition and predation), local environmental conditions, and substrate submersion time and depth (Chalmer, 1982; Bram et al., 2005). Competition among sessile fouling organisms can be through exploitation (competing for food) or interference (crushing, overgrowing, poisoning or smothering) (Arakawa, 1990; Lesser et al., 1992; Dalby and Young, 1993). The most common form of interference competition within fouling communities is overgrowing, a method used commonly by colonial ascidians and encrusting bryozoans to out-compete basal-growing individuals (Nandakumar, 1993). Fouling organisms also compete with the cultured bivalves for space, largely through overcrowding, which inhibits the space available for bivalve growth (Arakawa, 1990; Lesser et al., 1992). For bivalves, the strongest competitors for space are barnacles, ascidians, polychaetes and mussels (Arakawa, 1990). Competition for food between sessile filter-feeding organisms may occur because the availability of food is limiting in quantity and quality, depending on the spatial and temporal location of the community (Ward et al., 1998), overlapping diets and comparable filtration rates (dependent on species size and abundance) of the competing species (Buss and Jackson, 1981). Some organisms, such as bivalves, have the ability to adjust to food shortages by sorting and ingesting only particles with nutritional value; however, most filter-feeders (e.g. ascidians and some crustaceans) do not have the ability to sort particles in this way (Ward et al., 1998). Mytilus spp. and solitary ascidians, which have high feeding rates for particles of many types and sizes (Fiala-Medioni, 1978; Lesser et al., 1992), may be strong competitors for food with the cultured oysters. Predation is known to play a very important role in structuring marine hard-substrate communities (Dean and Hurd, 1980; Russ, 1980; Himmelman et al., 1983; Ambrose, 1991; Miron et al., 2002). Predation within a community provides ongoing disruption to community dynamics and creates patches of available substrate in an established community. Consumer disturbance can be the result of herbivores or carnivores (Wahl et al., 1997). Predation by herbivores or grazers on algae can enhance settlement of invertebrates by creating available space, or on the contrary, reduce settlement of invertebrates by the inadvertent ingestion of newly settled individuals (Dayton, 1971; Underwood et al., 1983). Ascidians may also act as predators by accidentally consuming invertebrate larvae while actively filter-feeding (Cowden et al., 1984;  5  Bingham and Walters, 1989). Fish have been documented as important predators in both intertidal and subtidal hard-substrate communities (Russ, 1980; Caine, 1989; Okamura, 1996; Connell and Anderson, 1999). Predation is a major concern for bivalve aquaculture. Cultured species, especially at young stages, are vulnerable to mobile predators such as drills or sea stars. Shellfish farmers in Prince Edward Island, Canada, identified the common sea star (Asterias vulgaris), moon shells (Lunatia heros) and the green crab (Carcinus maenas) as successful predators of cultured M. edulis and C. virginica (Miron et al., 2002). Shellfish farmers in BC have described predation by the common sea stars Evasterias troschelli and Pisaster ochraceus on suspended C. gigas. The presence of non-native species may play an important role in determining the structure of fouling communities. While many non-native species introduced into the marine environment have had very little effect on local community structure, a number of exotic species have completely altered community composition by out-competing native species for space and food to become the dominate species (Carlton, 1996; Kareiva, 1996; Cohen and Carlton, 1998; Stachowicz et al., 2002; Anil, 2006; Altman, 2007). Whether an introduced species becomes successfully established in a new environment depends on a number of environmental (water and current conditions) and community (species diversity) factors. The life history traits of the species, particularly its ability to compete, avoid predation and withstand variable water conditions, also play a significant role in the establishment of the species (Stachowicz and Whitlatch, 1999; Levine, 2000). The role played by biodiversity in hard-substrate communities is argued to be of critical importance in ensuring the proper functioning of the ecosystem and the ability of a community to resist the establishment of non-native species (Stachowicz and Whitlatch, 1999; Loreau et al., 2001). Introduced species may be so successful that they pose a threat to marine biodiversity, ecosystem function and, often, to the growth and health of economically important species (Travis, 1993; Stachowicz and Whitlatch, 1999; Holland, 2000; Covich et al., 2004; Anil, 2006; Thebault and Loreau, 2006). Introduced species in fouling communities that are reportedly important include: crustaceans Caprella mutica and Carcinus maenus (Grosholz, 2000; Anil, 2006; Ashton et al., 2007; Ashton et al., 2008), and ascidians Styela clava, Botrylloides violaceus, Botryllus schlosseri and Didemnum sp. (Grosberg, 1988; Stachowicz et al., 2002; Agius, 2007; Bourque, 2007; Bullard et al., 2007; Dijkistra et al., 2007b; Locke et al., 2007; McCarthy et al.,  6  2007; Osman and Whitlatch, 2007; Rajbanshi and Pederson, 2007; Valentine, 2007; Epelbaum et al., 2009). The importance of environmental conditions in determining marine invertebrate community composition cannot be overlooked because parameters, such as water column temperature and salinity, have significant effects on the settlement and recruitment of organisms. These conditions are difficult to study in a community context and thus are rarely discussed (Mazouni et al., 2001; Moura et al., 2007). A study on the interactions between the water column in a French lagoon and C. gigas in suspended culture, with the associated fouling community, discovered strong seasonal changes in the fouling community composition which were correlated with changes in the water parameters including temperature and nutrients (Mazouni et al., 2001). In the spring, the lagoon had high species diversity in the oyster fouling communities and low nutrient levels in the water column (except for nitrogen) while the autumn was characterized by low species diversity in the oyster fouling communities and high nutrient levels in the water column (Mazouni et al., 2001).  1.1.3  Treating fouling associated with bivalve aquaculture  The treatment, or removal, of fouling is costly to farmers (Enright, 1993; Guenther et al., 2006; Rodriguez and Ibarra-Obando, 2008). Antifouling studies have focused largely on mechanical and chemical treatments to control fouling. Mechanical treatments include burning (torch), drying (sun), boiling (water bath), removing (manual) or washing (high pressure) (Copisarow, 1945; Arakawa, 1990; Enright, 1993). In many cases, mechanical treatments have proved effective in managing fouling but they are generally too labour intensive to be cost-effective. For example, manual removal can account for more than 30% of the farm’s total operational costs (Claereboudt et al., 1994). Chemical treatments include dipping in brine, hydrated lime or acetic acid (Arakawa, 1990; Locke et al., 2009), or pre-treating bivalves and equipment before deployment. In the past, pre-treatments included toxic chemicals such as DDT, copper based paints or silicone coatings (Copisarow, 1945; Fry, 1975; Huguenin and Huguenin, 1982; Arakawa, 1990; Enright, 1993) and more recently non-toxic substances, which have not been named (Swain and Schultz, 1996; Swain et al., 1998). Chemical treatments, although widely used, particularly in the past, can have environmental implications due to the toxicity of the products used (Fry, 1975; Arakawa, 1990; Guenther et al., 2006). More recent efforts in controlling fouling have been directed towards biological treatments, the use of natural predators.  7  These include the use of sea urchins (Echinometra lucunter and Lytechinus variegates) on pearl oysters (Pinctada imbricata) in pearl nets (Lodeiros and Garcia, 2004); sea urchins (Echinus esculentus and Psammechinus miliaris) and hermit crabs (Pagurus sp.) on great scallops (Pecten maximus) in pearl nets (Ross et al., 2004); rock crab (Cancer irroratus) and hermit crab (Pagurus acadianus) on European oysters (Ostrea edulis) in lantern nets (Hidu et al., 1981; Enright et al., 1993); and dog whelks (Nucella lapillus) on European oysters (O. edulis) and great scallops (P. maximus) in trays (Minchin and Duggan, 1989). Despite the variety in cultured bivalve species and biological control species, all of the biological control studies listed above found a significant reduction in fouling and/or bivalve mortality with the introduction of a biological control. The use of biological controls in managing fouling communities associated with bivalve culture is clearly promising but more research is required for additional species of cultured bivalves and for bivalve farms in other regions. For example, there are no studies to date on either the use of biological controls on fouling communities on C. gigas or on any cultured bivalve farms in BC.  1.2  THESIS OVERVIEW  The overall research goal of my study was to examine the seasonal variation in the invertebrate fouling community associated with 2 different tray types (substrates) used at a deep-water suspended oyster farm in BC, Canada. The 2 main objectives of my study were to closely examine fouling community composition associated with a) the oyster trays (artificial/abiotic substrate) and b) the oysters (biotic substrate). Tray and oyster fouling communities were examined seasonally and between the 2 tray types. To achieve both objectives, the fouling community composition for both tray types was analyzed, every 3 months (excluding oyster fouling in the third 3-month interval) over the course of 1 year, to determine seasonal variation in invertebrate fouling. To assist in explaining seasonal changes in fouling community composition, water column parameters (temperature, pH, salinity, dissolved oxygen, chlorophyll and turbidity) were monitored on each sampling date.  8  LITERATURE CITED Agius, B.P., 2007. Spatial and temporal effects of pre-seeding plates with invasive ascidians. Journal of Experimental Marine Biology and Ecology 342: 30-39. Aleem, A.A., 1957. Succession of marine fouling organisms on test panels immersed in deepwater at la Jolla, California. Hydrobiologia 11: 40-58. Altman, S., Whitlatch, R.B., 2007. Effects of small-scale disturbance on invasion success in marine communities. Journal of Experimental Marine Biology and Ecology 342: 15-29. Ambrose, W.G., 1991. Are infaunal predators important in structuring soft-bottom communities? American Zoologist 31: 849-860. Anderson, M.J., 1999. Distinguishing direct from indirect effects of grazers in intertidal estuarine assemblages. Journal of Experimental Marine Biology and Ecology 234: 199-218. Anderson, M.J., Underwood, A.J., 1994. Effects of substratum on the recruitment and development of an intertidal estuarine fouling assemblage. Journal of Experimental Marine Biology and Ecology 184: 217-236. Anil, A.C., 2006. A perspective of marine bioinvasion. In multiple dimensions of global environmental change, TERI Press, India. Arakawa, K.Y., 1990. Competitors and fouling organisms in the hanging culture of the Pacific oyster, Crassostrea gigas (Thundberg). Marine and Freshwater Behaviour and Physiology 17: 67-94. Ashton, G.V., Willis, K.J., Cook, E.J., Burrows, M., 2007. Distribution of the introduced amphipod, Caprella mutica Schurin, 1935 (Amphipoda: Caprellida: Caprellidae) on the west coast of Scotland and a review of its global distribution. Hydrobiologia 590: 31-41. Ashton, G.V., Riedlecker, E.I., Ruiz, G.M., 2008. First non-native crustacean established in coastal waters of Alaska. Aquatic Biology 3: 133-137. Bayne, B.L., 1965. Growth and the delay of metamorphosis of the larvae of Mytilus edulis (L.). Ophelia 2: 1-47. BCSGA, 2009. British Columbia Shellfish Growers Association: Industry Encyclopedia. http://www.bcsga.ca/encyclopedia_index.php. Bonar, D.B., Weiner, R.M., Colwell, R.R., 1986. Microbial-invertebrate interactions and potential for biotechnology. Microbial Ecology 12: 101-110. Bourget, E., DeGuise, J., Daigle, G., 1994. Scales of substratum heterogeneity, substratum complexity and the early establishment of a marine epibenthic community. Journal of Experimental Marine Biology and Ecology 181: 31-51.  9  Bingham, B.L., Walters, L.J., 1989. Solitary ascidians as predators of invertebrate larvae: Evidence from gut analyses and plankton samples. Journal of Experimental Marine Biology and Ecology 131: 147-159. Bourque, D., Davidson, J., MacNair, N.G., Arsenault, G., LeBlanc, A.R., Landry, T., Miron, G., 2007. Reproduction and early life history of an invasive ascidian Styela clava Herdman in Prince Edward Island, Canada. Journal of Experimental Marine Biology and Ecology 342: 78-84. Bram, J.B., Page, H.M., Dugan, J.E., 2005. Spatial and temporal variability in early successional patterns of an invertebrate assemblage at an offshore oil platform. Journal of Experimental Marine Biology and Ecology 317: 223-237. Brown, K.M., Swearingen, D.C., 1998. Effects of seasonality, length of immersion, locality and predation on an intertidal fouling assemblage in the Northern Gulf of Mexico. Journal of Experimental Marine Biology and Ecology 225: 107-121. Bullard, S.G., Whitlatch, R.B., Osman, R.W., 2004. Checking the landing zone: Do invertebrate larvae avoid settling near superior spatial competitors? Marine Ecology Progress Series 280: 239-247. Bullard, S.G., Lambert, G., Carman, M.R., Byrnes, J., Whitlatch,, R.B., Ruiz, G., Miller, R.J., Harris, L., Valentine, P.C., Collie, J.S., Pederson, J., McNaught, D.C., Cohen, A.N., Asch, R.G., Dijkstra, J., Heinonen, K., 2007. The colonial ascidian Didemnum sp. A: Current distribution, basic biology and potential threat to marine communities of the northeast and west coast of North America. Journal of Experimental Marine Biology and Ecology 342: 99-108. Buss, L.W., Jackson, J.B.C., 1981. Planktonic food availability and suspension-feeder abundance: Evidence of in situ depletion. Journal of Experimental Marine Biology and Ecology 49: 151-161. Callow, M.E., Fletcher, R.L., 1994. The influence of low surface energy materials on bioadhesion- A review. International Biodeterioration and Biodegradation 34: 333-348. Caine, E.A., 1989. Caprellid amphipod behaviour and predatory strikes by fish. Journal of Experimental Marine Biology and Ecology 126: 173-180. Carlton, J.T., 1996. Pattern, process, and prediction in marine invasion ecology. Biological Conservation 78: 97-106. Chalmer, P.N., 1982. Settlement patterns of species in a marine fouling community and some mechanisms of succession. Journal of Experimental Marine Biology and Ecology 58: 7385. Claereboudt, M.R., Bureau, D., Côté, J., Himmelman, J.H., 1994. Fouling development and its effect on the growth of juvenile giant scallops (Placopecten magellanicus) in suspended culture. Aquaculture 121: 327-342.  10  Cohen, A.N., Carlton, J.T., 1998. Accelerating invasion rate in a highly invaded estuary. Science 279: 555-558. Connell, S.D., Anderson, M.J., 1999. Predation by fish on assemblages of intertidal epibiota: Effects of predator size and patch size. Journal of Experimental Marine Biology and Ecology 241: 15-29. Copisarow, M., 1945. Marine fouling and its prevention. Science 101: 406-407. Covich, A.P., Austen, M.C., Barlocher, F., Chauvet, E., Cardinale, B.J., Biles, C.L., Inchausti, P., Dangles, O., Solan, M., Gessner, M.O.., Statzner, B., Moss, B., 2004. The role of biodiversity in the functioning of freshwater and marine ecosystems. BioScience 54: 767775. Cowden, C., Young, C.M., Chia, F.S., 1984. Differential predation on marine invertebrate larvae by two benthic predators. Marine Ecology Progress Series 14: 145-149. Dahlem, C., Moran, P.J., Grants, T.R., 1984. Larval settlement of marine sessile invertebrates on surfaces of different colour and position. Ocean Science and Engineering 9: 225–236. Dalby, J.E., Young, C.M., 1993. Variable effects of ascidian competitors on oysters in a Florida epifaunal community. Journal of Experimental Marine Biology and Ecology 167: 47-57. Dayton, P.K., 1971. Competition, disturbance, and community organization: The provision and subsequent utilization of space in a rocky intertidal community. Ecological Monographs 41: 351–389. Dean, T.A., Hurd, L.E., 1980. Development in an estuarine fouling community: The influence of early colonists on later arrivals. Oecologia 46: 295-301. DFO, 2006. Assessing Habitat Risks Associated with Bivalve Aquaculture in the Marine Environment. DFO, Canadian Science Advisory Secretariat, Science Advisory Report 2006/005. DFO, 2008. Department of Fisheries and Oceans: Aquaculture Statistical Services. http://www.dfo-mpo.gc.ca/communic/statistics/aqua/aqua07_e.htm. Dijkstra, J., Harris, L.G., Westerman, E., 2007. Distribution and long-term temporal patterns of four invasive colonial ascidians in the Gulf of Maine. Journal of Experimental Marine Biology and Ecology 342: 61-68. Enright, C., 1993. Control of fouling in bivalve aquaculture. World Aquaculture 24: 44-46. Enright, C., Elner, R.W., Griswold, A., Borgese, E.M., 1993. Evaluation of crabs as control agents for biofouling in suspended culture of European oysters. World Aquaculture 24: 4951.  11  Epelbaum, A., Herborg, L.M., Therriault, T.W., Pearce, C.M., 2009. Temperature and salinity effects on growth, survival, reproduction, and potential distribution of two non-indigenous botryllid ascidians in British Columbia. Journal of Experimental Marine Biology and Ecology 369: 43-52. Erbland, P.J., Ozbay, G., 2008. A comparison of the macrofaunal communities inhabiting a Crassostrea virginica oyster reef and oyster aquaculture gear in Indian River Bay, Delaware. Journal of Shellfish Research 27: 757-768. Fiala-Medioni, A., 1978. Filter-feeding ethology of benthic invertebrates (Ascidians). IV. Pumping rate, filtration rate, filtration efficiency. Marine Biology 48: 243-249. Fry, W.G., 1975. Raft fouling in the Menai Strait, 1963-1971. Hydrobiologia 47: 527-558. Grosberg, R.K., 1988. Life-history variation within a population of the colonial ascidian Botryllus schlosseri. I. The genetic and environmental control of seasonal variation. Evolution 42: 900-920. Grosholz, E.D., Ruiz, G.M., Dean, C.A., Shirley, K.A., Maron, J.L., Connors, P.G., 2000. The impacts of a nonindigenous marine predator in a California Bay. Ecology 81: 1206-1224. Guenther, J., Southgate, P.C., de Nys, R., 2006. The effect of shell size on accumulation of fouling organisms on the Akoya pearl oyster Pinctada fucata (Gould). Aquaculture 253: 366-373. Gutiérrez, J.L., Jones, C.G., Strayer, D.L., Iribarne, O.O., 2003. Mollusks as ecosystem engineers: The role of shell production in aquatic habitats. Oikos 101: 79-90. Hidu, H., Conary, C., Chapman, S.R., 1981. Suspended culture of oysters: Biological fouling control. Aquaculture 22: 189-192. Himmelman, J.H., Cardinal, A., Bourget, E., 1983. Community development following removal of urchins, Strongylocentrotus droebachiensis, from the rocky subtidal zone of the St. Lawrence Estuary, eastern Canada. Oecologia 59: 27–39. Holland, B.S., 2000. Genetics of marine bioinvasions. Hydrobiologia 420: 63-71. Huguenin, J.E., Huguenin, S.S., 1982. Biofouling resistant shellfish trays. Journal of Shellfish Research 2: 41-46. Kareiva, P., 1996. Developing a predictive ecology for non-indigenous species and ecological invasions. Ecology 77: 1651-1652. LeBlanc, A.R., Landry, T., Miron, G. 2003. Identification of fouling organisms covering mussel lines and impacts of a common defouling method on the abundance of foulers in Tracadie Bay, Prince Edward Island. Canadian Technical Report of Fisheries and Aquatic Sciences 2477: 1-26.  12  Lesser, M.P., Shumway, S.E., Cucci, T., Smith, J., 1992. Impact of fouling organisms on mussel rope culture: Interspecific competition for food among suspension-feeding invertebrates. Journal of Experimental Marine Biology and Ecology 165: 91-102. Levine, J.M., 2000. Species diversity and biological invasions: Relating local process to community pattern. Science 288: 852-854. Locke, A., Doe, K.G., Fairchild, W.L., Jackman, P.M., Reese, E.J., 2009. Preliminary evaluation of effects of invasive tunicate management with acetic acid and calcium hydroxide on nontarget marine organisms in Prince Edward Island, Canada. Aquatic Invasions 4: 221-236. Locke, A., Hanson, J.M., Ellis, K.M., 2007. Invasion of the southern Gulf of St. Lawerence by the clubbed tunicate (Styela clava Herdman): Potential mechanisms for invasions of Prince Edward Island estuaries. Journal of Experimental Marine Biology and Ecology 342: 69-77. Lodeiros, C., Garcia, N., 2004. The use of sea urchins to control fouling during suspended culture of bivalves. Aquaculture 231: 293-298. Lodeiros, C.J.M., Himmelman, J.H., 2000. Identification of factors affecting growth and survival of the tropical scallop Euvola (Pecten) ziczac in the Golfo de Cariaco, Venezuela. Aquaculture 182: 91-114. Loreau, M., Naeem, S., Inchausti, P., Bengtsson, J., Grime, J.P., Hector, A., Hooper, D.U., Huston, M.A., Raffaelli, D., Schmid, B., Tilman, D., Wardle, D.A., 2001. Biodiversity and ecosystem functioning: Current knowledge and future challenges. Science 294: 804-808. Mazouni, N., Gaertner, J.C., Deslous-Paoli, J.M., 2001. Composition of biofouling communities on suspended oyster cultures: An in situ study of their interactions with the water column. Marine Ecology Progress Series 214: 93-102. McCarthy, A., Osman, R.W., Whitlatch, R.B., 2007. Effects of temperature on growth rates of colonial ascidians: A comparison of Didemnum sp. to Botryllus schlosseri and Botrylloides violaceus. Journal of Experimental Marine Biology and Ecology 342: 172-174. McKindsey, C.W., Anderson, M.R., Barnes, P., Courtenay, S., Landry, T., Skinner, M., 2006. Effects of shellfish aquaculture on fish habitat. DFO, Canadian Science Advisory Secretariat, Research Document 2006/011. Minchin, D., Duggan, C.B., 1989. Biological control of the mussel in shellfish culture. Aquaculture 81: 97-100. Miron, G., Landry, T., MacNair, N., 2002. Predation potential by various epibenthic organisms on commercial bivalve species in Prince Edward Island: Preliminary results. Canadian Technical Report of Fisheries and Aquatic Sciences 2392: 1-44. Moura, A., Boaventura, D., Curdia, J., Carvalho, S., Cancela de Fonseca, L., Leitao, F.M., Santos, M.N., Monteiro, C.C., 2007. Effect of depth and reef structure on early macrobenthic  13  communities of the Algrave artificial reefs (southern Portugal). Hydrobiologia 580: 173180. Murad, B.M., Mohammad, F.L.S., 1976. Relationship between biofouling and growth of the pearl oyster Pinctada fucata (Gould) in Kuwiat, Arabian Gulf. Hydrobiologia 51: 129-138. Nandakumar, K.U., 1993. Interspecific competition among fouling organisms in Tomioka Bay, Japan. Marine Ecology Progress Series 94: 43-50. Okamura, B., 1986. Formation and disruption of aggregations of Mytilus edulis in the fouling community of San Francisco Bay, California. Marine Ecology Progress Series 38: 275-282. Osman, R.W., 1977. The establishment and development of a marine epifaunal community. Ecological Monographs 47: 37-63. Osman, R.W., Whitlatch, R.B., 2007. Variation in the ability of Didemnum sp. to invade established communities. Journal of Experimental Marine Biology and Ecology 342: 40-53. Parsons, G.J., Dadswell, M.J., 1992. Effect of stocking density on growth, production, and survival of the giant scallop, Placopecten magellanicus, held in intermediate suspension culture in Passamaquoddy Bay, New Brunswick. Aquaculture 103: 291-309. Qian, P-Y., 1999. Larval settlement of polychaetes. Hydrobiologia 402: 239-253. Railkin, A.I., 2004. Marine biofouling: Colonization processes and defenses. CRC Press, Boca Raton, Florida. Rajbanshi, R., Pederson, J., 2007. Competition among invading ascidians and a native mussel. Journal of Experimental Marine Biology and Ecology 342: 163-165. Rodriguez, L.F., Ibarra-Obando, S.E., 2008. Cover and colonization of commercial oyster (Crassostrea gigas) shells by fouling organisms in San Quintin, Mexico. Journal of Shellfish Research 27: 337-343. Ross, K.A., Thorpe, J.P., Brand, A.R., 2004. Biological control of fouling in suspended scallop cultivation. Aquaculture 229: 99-116. Russ, G.R., 1980. Effects of predation by fishes, competition, and structural complexity of the substratum on the establishment of a marine epifaunal community. Journal of Experimental Marine Biology and Ecology 42: 55-69. Stachowicz, J.J., Whitlatch, R.B., 1999. Species diversity and invasion resistance in a marine ecosystem. Science 286: 1577-1580. Stachowicz, J.J., Fried, H., Osman, R.W., Whitlatch, R.B., 2002. Biodiversity, invasion resistance and marine ecosystem function: Reconciling pattern and process. Ecology 83: 2575-2590.  14  Swain, G.W.J, Schultz, M.P., 1996. The testing and evaluation of non-toxic antifouling coatings. Biofouling 10: 187-197. Swain, G.W., Nelson, W.G., Preedeekanit, S., 1998. The influence of biofouling adhesion and biotic disturbance on the development of fouling communities on non-toxic surfaces. Biofouling 12: 257-269. Taylor, J.J., Southgate, P.C., Rose, R.A., 1997. Fouling animals and their effect on the growth of silver-lip pearl oysters, Pinctada maxima (Jameson) in suspended culture. Aquaculture 153: 31-40. Thébault, E., Loreau, M., 2006. The relationship between biodiversity and ecosystem functioning in food webs. Ecological Research 21: 17-25. Travis, J., 1993. Invader threatens Black, Azov Seas. Science 262: 1366-1367. Tyrrell, M.C., Byers, J.E., 2007. Do artificial substrates favour nonindigenous fouling species over native species? Journal of Experimental Marine Biology and Ecology 342: 54-60. Underwood, A.J., Chapman, M.G., 2006. Early development of subtidal macrofaunal assemblages: Relationships to period and timing of colonization. Journal of Experimental Marine Biology and Ecology 330: 221-233. Underwood, A.J., Denley, E.J., Moran, M.J., 1983. Experimental analyses of the structure and dynamics of mid-shore rocky intertidal communities in New South Wales. Oecologia 56: 202–219. Valentine, P.C., Carman, M.R., Blackwood, D.S., Heffron, E.J., 2007. Ecological observations on the colonial ascidian Didemnum sp. in a New England tide pool habitat. Journal of Experimental Marine Biology and Ecology 342: 109-121. Wahl, M., 1989. Marine epibiosis. I. Fouling and biofouling: Some basic aspects. Marine Ecology Progress Series 58: 175-189. Wahl, M., Hay, M.E., Enderlein, P., 1997. Effects of epibiosis on consumer-prey interactions. Hydrobiologia 355: 49-59. Walters, L.J., 1992. Post-settlement success of the absorbent bryozoan Bugula neritina (L.): the importance of structural complexity. Journal of Experimental Marine Biology and Ecology 164: 55-71. Walters, L.J., Wethey, D.S., 1991. Settlement, refugees and adult body form in colonial marine invertebrates: A field experiment. Biological Bulletin 180: 112-118. Ward, J.E., Levinton, J.S., Shumway, S.E., Cucci, T., 1998. Particle sorting in bivalves: In vivo determination of the pallial organs of selection. Marine Biology 131: 283-292.  15  Woodin, S.A., 1991. Recruitment of infauna: Positive or negative cues. American Zoologist 31: 797-807.  16  CHAPTER TWO. Effects of season and tray type on fouling communities associated with oyster trays 1 2.1  INTRODUCTION  Fouling, or the colonization of a substrate by unwanted marine plants and animals, in association with deep-water bivalve aquaculture is of increasing concern to shellfish growers. Fouling communities growing on such facilities can lead to reduced growth and survival of the cultured species, increased weight and drag on the equipment and increased labour costs associated with treating or removing fouling (Enright, 1993; Guenther et al., 2006; Rodriguez and Ibarra-Obando, 2008). The introduction of a suspended bivalve facility into the marine environment provides a three-dimensional structure with an abundance of abiotic (equipment) and biotic (bivalve) substrates suitable for the settlement and growth of marine organisms (DFO, 2006). It has been well-documented that fouling community composition varies spatially and temporally and is strongly influenced by factors such as larval recruitment, substrate type, disturbances such as competition and predation, local environmental conditions and speciesspecific traits (Dean and Hurd, 1980; Chalmer, 1982; Ward et al., 1998; Miron et al., 2002; Bram et al., 2005; Moura et al., 2007). There are numerous studies that have examined the local colonization of artificial substrates including glass, asbestos, cement, asbestos-cement, plastic, wood, ceramic/clay, and polyvinyl chloride (e.g. Aleem, 1957; Fry, 1975; Anderson and Underwood, 1994; Tyrrell and Byers, 2007), by suspending replicate panels of substrate in the water column and monitoring larval settlement and invertebrate growth over time. Much of the previous work focused on the change in dominant species in an attempt to document succession and does not examine changes in overall community composition. Moreover, these studies were highly variable (different locales, different depths, submersed at different times of year for different lengths of time) and generalizations are therefore extremely difficult. With that said, these studies do provide useful details of colonization for a few common groups, or species of invertebrates (e.g. Mytilus spp., ascidians, Balanus spp., hydroids and encrusting bryozoans) that occupied most panels at some stage in the colonization process. To date, there is a dearth of literature on the colonization and community composition of fouling communities associated with bivalve aquaculture.  1  A version of this chapter will be submitted for publication. Switzer, S.E., Barnes, P.A., McKinley, R.S. Effects of season and tray type on fouling communities associated with oyster trays.  17  In BC, oysters intended for the half-shell market are most often grown-out in trays suspended from rafts (BCSGA, 2009). There are a number of tray types used to grow oysters, all designed to allow space for oysters to grow and to provide adequate water flow. Recently in BC, a metal tray with polyvinyl coating has begun replacing the “traditional” plastic tray, with the aims of increasing oyster density and reducing fouling. When oyster trays become heavily fouled, the flow of water through the tray is restricted and as a result the food and oxygen available to the oysters is limited (Enright, 1993; Huguenin and Huguenin, 1982). Fouled trays must be cleaned regularly, resulting in increased labour costs. The purpose of this study was to examine seasonal variation in the composition of invertebrate fouling communities associated with 2 different tray types (“traditional” plastic and new polyvinyl) used at a deep-water suspended Pacific oyster farm in BC. The fouling communities on the seeded experimental trays were examined every 3 months over the course of a year. An understanding of the seasonal variation of fouling communities associated with the 2 tray types used in BC will assist oyster farmers in determining which tray type best reduces fouling as well as when, and what type of, antifouling measures might be appropriate.  2.2  METHODS  2.2.1  Study site and experimental design  The study was conducted at a deep-water suspended Pacific oyster (C. gigas) aquaculture farm located in Sansum Narrows, between Saltspring Island and Vancouver Island, BC, Canada (48°46.371' N, 123°32.984' W ') (Figure 2.1a). Sansum Narrows is a small passage with very little commercial vessel traffic or anthropogenic impacts (adjacent land is scarcely occupied) with strong currents at flood tide. The oyster farm is situated in a small embayment on the east side of the Narrows closest to Saltspring Island (Figure 2.1). The site is relatively small in size, supporting 9 standard aquaculture rafts (80 stacks of trays per raft), situated in groups of 3, and a large raft (120 stacks of trays) that also serves as a work and storage area. The experimental oyster raft (Figure 2.1b inset) used in this study was positioned in the southeastern corner of the embayment, on the north edge of a set of 3 rafts (Figure 2.1b). Of the 80 oyster stacks on the experimental raft, 74 stacks were under the supervision of the farm personnel while 3 stacks of “traditional” plastic trays (PL) and 3 stacks of the new metal tray with polyvinyl coating (PV) were used in this study (Figure 2.2a and 2.2b). The 2 tray types had the same overall dimensions but key differences included tray material, size of the holes in  18  the material (available surface area), presence or absence of a tray liner and “stack-ability” of the trays (Figure 2.2c and 2.2d). Refer to Table 2.1 for a detailed comparison.  a  b  NWF NWR  C SER  R  Figure 2.1. Location of oyster farm, experimental raft and water parameter stations. (a) Oyster farm ( ) (DFO, 2005), (b) Oyster farm in the embayment (photo by Google Earth), the experimental raft (black rectangle and inset photo) and the stations used in the collection of water column parameter data, 3 raft stations (NW corner (NWR), centre (C) and SE corner (SER)), a station within the farm (NW corner (NWF)) and a reference station (R).  a  b  c  d  Figure 2.2. Experimental oyster culture trays. (a) “traditional” plastic oyster trays stacked seven trays high (note gaps between trays), (b) polyvinyl oyster trays stacked ten trays high (notice trays fit tightly together), (c) plastic oyster tray (PL) and (d) new metal oyster tray with polyvinyl coating (PV).  19  Table 2.1. Comparison of key differences between the plastic (PL) and polyvinyl (PV) experimental trays. Plastic Tray (PL)  Polyvinyl Tray (PV)  Material Type  plastic  metal with polyvinyl coating  Holes in Material  variable-sized, large handles  uniform-sized, no handles  Liner Use  none  plastic with small holes, lines tray bottom  “Stack-ability”  7 trays per stack with gaps between trays  10 trays per stack, trays fit tightly with no gaps  All tray stacks on the experimental oyster raft were seeded in October 2006 following standard operating procedures used on the farm: an estimated 120 juvenile oysters, approximately 1 year old, were placed in each tray. Experimental oyster stacks were suspended from the raft in randomly-selected locations with the first tray in each stack positioned at 1.5m below the surface. Every 3 months for 12 months (January 2007, April 2007, July 2007 and October 2007), the experimental stacks were retrieved and the fouling organisms meticulously collected from the third tray (oyster fouling was removed and retained) in each experimental stack. Fouling was roughly cleaned from all other trays in the stack using a wire brush before redeployment.  2.2.2  Field methods  In January, April, July and October 2007, experimental stacks were pulled using the farm’s crane and the experimental tray in each stack was retrieved for processing. After removing the oysters from each of the 6 experimental trays, fouling was carefully collected from each tray using forceps, scalpels and a soft-wire brush. Fouling was placed into seawater-filled totes that were fitted with a spout and 500µm Nitex filter, to enable seawater replacement without loss of organisms. Totes were transported to the laboratory at the end of each collection day where they were immediately filtered and preserved (see section 2.2.3). Fouling was collected from the wrong experimental tray in 1 of the PL stacks in July 2007 resulting in only 2 fouling sample replicates for July PL-trays In October 2006, Stowaway Tidbit temperature loggers (Model No. TBI32-05+37; Onset Computer Corporation) were deployed at depths of 1m, 3m, 5m, 9m and 11m in the centre of the experimental raft. The loggers monitored water temperature every 15 minutes over the course of the study and, every sampling date, these data were uploaded in the field to an optic shuttle for return to the laboratory and the loggers were redeployed. On each sampling date, a YSI 660  20  Sonde (multi-parameter water quality monitor) with a 650 handheld display unit and a 30m cable, were used to collect water column profile data at 5 stations. Three of the stations were within the experimental raft (NW corner (NWR), centre (C) and SE corner (SER)), a fourth station was within the farm (NW corner (NWF)) and a fifth (reference) station (R) was located southeast of the farm in the Narrows (Figure 2.1b). Temperature, pH, chlorophyll, turbidity, salinity and dissolved oxygen were measured using the Sonde at each station at every meter of depth to a maximum of 30m or until reaching bottom. Turbidity and chlorophyll measurements were ground-truthed in the laboratory by analyzing water samples collected using a 2-litre Niskin water sampler positioned alongside the Sonde at 3m, 5m and 11m at each of the 5 stations.  2.2.3  Laboratory methods  The fouling samples were filtered immediately upon arrival at the Centre for Shellfish Research (Vancouver Island University), using a 500µm screen and then preserved in 10% buffered formalin for 3 days before being transferred to 75% ethanol for long term storage. Organisms in the preserved samples were separated from the organic matter using a dissecting microscope and then identified to species level or to the lowest taxon possible. The number of individuals of each species was recorded and each individual was placed within a size category based on the known adult size for that species. Representatives of each size category for each species were weighed (wet weights) for biomass estimates. Because the fouling samples proved time consuming to process (e.g. 50 hours/sample for the seasonally-depauperate January samples), a subsampling method was devised, tested and then introduced to reduce the processing time. The subsampling method required removal and identification of the large organisms, followed by a fixed-fraction split on the remaining sample (Barbour and Gerritsen, 1998). Fouling samples from January, April and October 2007 were processed using a 50% fixed-fraction split. As the July fouling samples were up to 3 times the volume of the samples in January, an alternative processing method was required. Initially, 3 of the 5 July samples were processed using a 50% fixed-fraction split on the whole sample. In the half-sample chosen for processing, crustaceans (which were visibly the most abundant taxon) were separated from all other organisms. The crustacean sample then underwent an additional split into eighths (such that the final crustacean subsample used for identification was a 6.2% fraction) while all other organisms were sorted and identified from the 50% fraction subsample. Despite this reduction in sample size, >200 hours was still required to process a single July  21  subsample and another alternative method had to be introduced to process the remaining 2 July samples. The 2 remaining July samples were split into sixteenths (6.2% fixed-fraction); however, 3 of the 6.2% fractions were processed to ensure minimal loss of species for a total fraction of 18.6%. All fouling data from the fractionated subsamples were multiplied to determine totals for the whole samples. Tidbit temperature data was downloaded from the optic shuttle to a computer and tabulated in preparation for analysis. Water column data collected in the field using the Sonde profiler was downloaded and used to generate seasonal water column profiles for the 6 parameters (temperature, pH, chlorophyll, turbidity, salinity and dissolved oxygen) at each of the 5 stations. Niskin water samples were frozen (-4°C) immediately after returning from the field until 2 days prior to analysis when they were placed in a dark fridge to thaw. Samples were analyzed in November 2007, a maximum of 13 months after collection. Chlorophyll a, b and c were extracted from Niskin seawater samples using acetone and then measured using a Biochrom Ultraspec 2100pr UV/Visible spectrophotometer (Arar, 1997). Measures of chlorophyll a, b and c were then combined to estimate total chlorophyll. Determination of total suspended solids (TSS) followed the method of Moran et al. (1999). The seawater samples were passed through a pre-weighed glass fiber filter, then the filter and collected solids were dried and weighed. Because Sonde chlorophyll and turbidity probes are known to have poor accuracy at very low levels (~0µg/L and ~0NTU) (YSI, pers. comm.), profiles of chlorophyll and total suspended solids, created with data collected from processing the Niskin water samples, were compared with the Sonde chlorophyll and turbidity profiles to ensure accuracy of the Sonde chlorophyll and turbidity probes. The Sonde levels were much lower than those measured on the water samples and therefore, the Sonde data for these 2 variables will not be presented or discussed further herein.  2.2.4  Data analysis  The number of replicates of oyster and tray fouling community samples are shown in Table 2.2. The fouling community composition was summarized for: abundance (total number of individuals per tray and number of individuals per major taxonomic group per tray), biomass (total organismal weight per tray and organismal weight per major taxonomic group per tray) and species richness (total number of species per tray and number of species per major taxonomic group per tray). For invertebrates, “major taxonomic group” refers to class: invertebrate species were grouped according to class and the data combined. Non-invertebrates present, Foraminifera  22  and Actinopterygii, were also categorized as “major taxonomic groups” for the purpose of data analyses. Although data were collected for individual species, this dataset was used only in multivariate analyses. Community dominance was calculated for each tray sample using the Swartz Dominance Index (SDI). SDI consists of the minimum number of species that make up 75% of the total abundance (Swartz, 1978). Univariate analyses were carried out using NCSS statistical software (Hintze, 2007) with a rejection criterion of α=0.050. Abundance, biomass and species richness (total and for each major group) and SDI values were compared between dates and tray types using a Two-Factor Analysis of Variance ANOVA), followed by post-hoc multiple comparison Tukey Tests. Normality of data was tested using a number of tests, including the Shapiro-Wilk, the Anderson-Darling, D’Agostino Skewness and D’Agostino Kurtosis. To satisfy the assumption of normality for the parametric ANOVA and Tukey tests, all abundance and biomass data were log-transformed. In this study, results of the ANOVA and the Tukey post-hoc tests occasionally conflicted with each other. In most of these cases, the ANOVA detected a significant difference that was not reflected by the Tukey tests. Conflicts like these occur because the ANOVA utilizes pooled data for all comparisons, thus having a higher sample size and degrees of freedom, and a lower overall variance than the Tukey tests. For consistency, ANOVA results are reported and when supported by Tukey tests, these results are discussed. Table 2.2. Number of replicate samples for each fouling community (date and tray type). Sampling Date  Plastic Tray (PL)  Polyvinyl Tray (PV)  January 2007  3  3  April 2007  3  3  July 2007  2*  3  October 2007  3  3  * Note 2 replicate samples in July due to a mistake made in sample collection.  Multivariate analyses were conducted using the Bray-Curtis dissimilarity coefficient (Bray and Curtis, 1957) for pair-wise comparisons of species abundance and species biomass. The pattern of dissimilarities between all paired samples was then plotted using a hierarchical cluster dendrogram and an unweighted pair-group mean average sort algorithm (Sneath and Sokal, 1973). To objectively analyze the between-group versus the within-group variance at each cluster linkage, and thereby determine whether the communities linked were representative of the same community, a non-parametric bootstrap method (SIGTREE) was used to test the null hypothesis  23  that the 2 groups being linked together at any given linkage were the same (α=0.050). The bootstrap method was run using 200 simulations for each analysis (Nemec and Brinkhurst, 1988).  2.3  RESULTS  2.3.1  Composition of fouling communities on oyster-culture trays  Organisms in the fouling community samples collected from the 2 tray types in January, April, July and October 2007 belonged to 22 major taxonomic groups: 20 invertebrate classes and the non-invertebrate Phyla Foraminifera and Class Actinopterygii (Table 2.3). In total, the samples contained 1,311,637 individuals belonging to 182 invertebrate species, 10 Foraminifera species and 2 species of Actinopterygii (fish), with a total biomass of all fouling individuals of 2.292kg. Univariate analyses of abundance of individuals (total and per major taxonomic group), species richness (total and per major taxonomic group), biomass (total and per major taxonomic group) and a measure of dominance (Swartz Dominance Index), as well as multivariate analysis of species abundance and species biomass, were used to compare fouling communities between sampling dates and tray types. 2.3.1.1 Abundance The total abundance of individuals (log transformed) in the tray fouling communities was different between sampling dates (data pooled across tray type), but not between tray types (data pooled across date) (Table 2.4). For both tray types, the July communities had the highest total abundances of all dates (PL-tray 314461+56715indiv/tray; PV-tray 181789+106840indiv/tray) and January communities had the lowest abundances (PL-tray 474+77indiv/tray; PV-tray 1399+934indiv/tray) (Figure 2.3a). For both tray types, the January and July communities were different both from each other and from the April and October communities (Tukey F=4.94, DF= 15, p<0.05); the April and October communities were not different from each other. Hence, the total abundance data shows a clear seasonal trend with high abundance in summer, low abundance in winter and spring and fall abundances similar to each other and falling between the summer and winter values. Tray type had no effect on the total abundances at any time. Abundance data (log transformed) for the 22 major groups, showed results similar to those for total abundance; for 18 of the 22 groups abundance was significantly different between sampling dates (Table 2.3). The abundance of most major groups was highest in July and lowest in January, with the abundance in April and October falling in between these extremes (for example, see Figure 2.4a). A notable exception to this pattern was the Class Demospongiae for  24  which abundance in the October communities from both tray types was significantly higher than abundance in the January PL-tray and April PV-tray communities, and was greater in the October PL-tray community than in the July PL-tray community (Tukey F=4.94, DF= 15, p<0.05) (Figure 2.4b). Other exceptions were Class Echinoidea, for which abundance in both tray types was greater in April than in January, and Class Turbellaria, for which abundance in both tray types was greater in April than all other sampling dates. Table 2.3. Summary of two-factor ANOVA results (p-values) for the 22 major taxonomic groups for abundance (log (x+0.1) transformed number individuals per tray), biomass (log (x+0.1) grams per tray) transformed and, species richness (number species per tray), for 2 factors (date and tray type) and the interaction between the 2 factors. * indicates a significant p-value, ** indicates a significant ANOVA (pooled data) result not supported by Tukey posthoc results (non-pooled data) Abundance Major Group Polychaeta Malacostraca Maxillapoda Ostracoda Pycnogonida Bivalvia Gastropoda Polyplacophora Asteroidea Echinoidea Holothuridea Ophiuroidea Ascidiacea Demospongiae Gymnolaemata Anopla Enopla Anthozoa Hydrozoa Turbellaria Foraminifera Actinopterygii  Biomass  Species Richness  Date  Tray Type  Interaction  Date  Tray Type  Interaction  Date  Tray Type  Interaction  <0.01 * <0.01 * <0.01 * <0.01 * <0.01 * <0.01 * <0.01 * <0.01 * <0.01 * <0.01 * <0.01 * 0.62 <0.01 * <0.01 * <0.01 * 0.45 0.02 ** 0.48 <0.01 * <0.01 * <0.01 * 0.69  <0.01 * 0.33 0.27 0.97 0.96 0.03 ** 0.02 ** 0.17 0.50 0.43 0.88 0.21 0.73 0.16 0.64 0.36 0.45 0.06 0.33 0.74 0.50 0.41  <0.01 * 0.09 0.54 0.32 0.07 0.13 0.44 0.27 0.34 0.88 0.12 0.62 0.55 0.09 0.71 0.45 0.84 0.86 0.29 0.29 0.27 0.24  <0.01 * <0.01 * <0.01 * 0.02 ** 0.10 <0.01 * 0.01 ** 0.16 <0.01 * <0.01 * 0.16 0.46 <0.01 * <0.01 * <0.01 * 0.45 <0.01 ** 0.28 <0.01 * <0.01 * <0.01 * 0.71  0.02 ** 0.83 0.09 0.88 0.74 0.32 0.29 0.47 0.04 ** 0.29 0.12 0.35 0.78 0.93 0.89 0.36 0.82 0.01 ** 0.58 0.89 0.42 0.55  0.07 0.49 0.11 0.99 0.99 0.54 0.26 0.41 0.61 0.78 0.40 0.46 0.22 0.90 0.37 0.45 0.97 0.29 0.40 0.76 0.79 0.24  <0.01 * <0.01 * NA <0.01 * <0.01 * <0.01 * <0.01 * <0.01 * 0.02 ** 0.02 ** <0.01 * 0.63 <0.01 ** <0.01 * <0.01 * 0.45 0.09 0.54 <0.01 * <0.01 * <0.01 * 0.69  <0.01 * 0.10 NA 1.00 1.00 0.02 ** 0.02 ** 0.53 0.23 0.27 0.12 0.20 0.64 0.11 0.22 0.36 0.73 0.10 0.41 0.36 <0.01 ** 0.41  0.01 * 0.02 * NA 0.73 0.02 * 0.29 0.21 0.24 0.59 0.94 0.34 0.63 0.64 0.01 * 0.17 0.45 0.70 0.71 0.60 0.45 0.02 * 0.24  Table 2.4. Summary of two-factor ANOVA results for total abundance (log (x+0.1) transformed total number of individuals per tray), biomass (log (x+0.1) transformed total biomass per tray) and species richness (total number of species per tray) for 2 factors (date and tray type) and for the interaction between the 2 factors. * indicates a significant p-value Abundance  Biomass  Species Richness  Date  Tray Type  Interaction  Date  Tray Type  Interaction  Date  Tray Type  Interaction  F-Ratio  139.22  2.63  2.59  37.05  0.20  0.61  52.32  16.91  5.04  p-value  <0.01 *  0.13  0.09  <0.01 *  0.66  0.62  <0.01 *  <0.01 *  0.01 *  Power (α=0.05)  1.00  0.33  0.52  1.00  0.07  0.15  1.00  0.97  0.87  25  b  c 3  Tray_Type PL PV  log_Total Biomass (g / tray)  log_Total Abundance (# indiv. / tray)  6  5  4  2  1  0  Total Species Richness (# species / tray)  a  Tray_Type PL PV  2  1  0  1 Jan n=3  2 Jul 3 Oct 4 Apr n=3Date n=2 n=3  1 Jan n=3  100  Tray_Type PL PV  80  60  40  20  0  2 Jul 3 Oct 4 Apr n=3Date n=2 n=3  1 Jan n=3  2 3 Oct 4 Apr Jul Date n=3 n=2 n=3  Figure 2.3. Seasonal changes in fouling communities on suspended PL and PV trays. (a) Mean total abundance (log transformed) per fouling community (number of individuals per tray + SE), (b) Mean total biomass (log transformed) per fouling community (g per tray + SE) and (c) Mean total species richness per fouling community (number of species per tray + SE).  6 5  Tray_Type PL PV  4 3 2 1 0 Jan 1  Apr 2  Jul 3  Oct 4  c  2 Tray_Type PL PV  1  0  -1 Jan 1  Apr 2  Date  Jul 3  Date  Oct 4  log_Polychaeta Abundance ( # indiv. / tray)  b log_Demospon. Abundance ( # indiv. / tray)  log_Bivalvia Abundance ( # indiv. / tray)  a  5 Tray_Type PL PV  4  3  2  1  0 Jan 1  Apr 2  Jul 3  Oct 4  Date  Figure 2.4. Seasonal mean abundance (log transformed) per tray fouling community (number individuals per tray) on suspended PL and PV trays for (a) Class Bivalvia, (b) Class Demospongiae and (c) Class Polychaeta.  The Class Polychaeta was the only major group with a difference in abundance between tray types, with higher abundance in the July PL-trays than in the July PV-trays (Tukey F=4.94, DF= 15, p<0.05) (Figure 2.4c). A significant interaction for Class Polychaeta suggested that season and tray type interact together to produce a higher Polychaeta abundance in July PL-trays. Although a significance level of 0.05 was chosen for these analyses, it is worth noting the  26  marginal significance of tray type for Anthozoa abundance and the interaction of date and tray type for Pycnogonida (Table 2.3). In summary, abundance trends within major groups were similar to total abundance trends, showing seasonal variation (summer highs and winter lows) and, with the exception of Polychaeta, a lack of significant variation between tray types. Total abundance for both tray types was dominated by Polychaeta in January and Malacostraca in April. In July, total abundance in PL-trays was dominated by Bivalvia followed by Malacostraca and in PV-trays by Malacostraca followed by Bivalvia. In October, total abundance was dominated by Polychaeta followed by Malacostraca in PL-trays and by Malacostraca followed by Polychaeta in PV-trays. When Class Polychaeta dominated, (January both tray types and October PL-tray), Platynereis bicanaliculata and Chrysopetalum occidentale were the most abundant polychaete species present. When Class Malacostraca dominated (April both tray types, July PV-tray and October PV-tray), Jassa staudei, Gammaridea sp. and Caprella mutica were the most abundant species. Mytilus sp. was the most abundant species when Class Bivalvia dominated (July PL-tray).  2.3.1.2 Biomass Fouling community total biomass demonstrated seasonal variation but no variation between the 2 tray types (Table 2.4). Figure 2.3b illustrates that for both tray types, total biomass peaked in July, followed by October, April and January. In fact, the total biomass of July communities in each of the tray types (PL-tray 144+314g/tray; PV-tray 335+228g/tray) was higher than the total biomass of the January and April communities in the corresponding tray type (Tukey F=4.90, DF= 15, p<0.05). In addition, the total biomass of October communities (both tray types) was higher than that of the January communities (Tukey F=4.90, DF= 15, p<0.05). In general, total biomass showed a seasonal trend similar to total abundance, with largest biomass in the summer (July) and smallest biomass in the winter (January). Unlike abundance, however, biomass was greater in the fall (October) when individual organisms were larger in size, than in the spring (April) when individuals were more abundant. Analysis indicated that tray type had no affect on either total abundance or total biomass on any of the sampling dates (Table 2.4). Biomass results for the 22 major groups indicated seasonal changes in 15 major groups and tray type differences in 3 major groups (Table 2.3). Similar to total biomass, seasonal changes in the biomass of most major groups were driven largely by the high biomass of the July communities and the low biomass of the January communities (for examples, see Figure 2.5a and  27  2.5b). Notable exceptions were the Asteroidea (Figure 2.5c), Echinoidea and Demospongiae, which had greater biomass in the October fouling communities (both tray types) than any other sampling date and the Turbellaria which had greater biomass in the April fouling communities (both tray types) than any other sampling date (all groups Tukey F=4.90, DF= 15, p<0.05). b  c 3  Tray_Type PL PV  log_Malacostraca Biomass ( g / tray)  log_Bivalvia Biomass ( g / tray)  3  2  1  0  1  Tray_Type PL PV  log_Asteroidea Biomass ( g / tray)  a  2  1  0  1 Jan  2 Apr  3Jul  Date  4 Oct  0  -1  -2  -1  -1  Tray_Type PL PV  1 Jan  2 Apr  3 Jul  4 Oct  1 Jan  2 Apr  3 Jul  4 Oct  Figure 2.5. Seasonal mean biomass (log transformation) per tray fouling community (g per tray + SE) on suspended PL and PV trays for (a) Class Bivalvia, (b) Class Malacostraca and (c) Class Asteroidea.  Class Polychaeta, Asteroidea and Anthozoa all had greater biomass in the PL-trays than the PV-trays (Table 2.3). While the higher biomass on the PL-trays for Class Polychaeta was supported by a higher abundance, the higher biomass for Asteroidea and Anthozoa was supported by the presence of larger individuals. There was no significant interaction for any of the major groups, however, Class Polychaeta had marginal significance (Table 2.3). Malacostraca made the greatest contribution to biomass in 5 of the 8 fouling communities sampled (April PL-tray and both tray types in July and October); Malacostraca biomass comprised between 43% and 59% of the total biomass of these communities. In the remaining 3 communities, the greatest contributors to total community biomass were the Polychaeta (January PL-tray), Actinopterygii (January PV-tray) and Gymnolaemata (April PV-tray). The Bivalvia (July), Gastropoda (January), Holothuridea (October), Ascidiacea (July) and Echinoidea (October) also made significant contributions to the total community biomass, depending on the sampling date. 2.3.1.3 Species richness The species richness data were normally distributed and did not require transformation. Species richness in the PL-tray fouling communities was highest in July, followed by April, October and  28  January respectively (Figure 2.3c). In contrast, species richness in the PV-tray fouling communities was highest in April, followed by October, July and January respectively. Total species richness was significantly different between dates and between tray types (Table 2.4). Tukey post-hoc tests revealed that total species richness for the PL-tray communities was significantly lower in January (53+7species/tray) than in the other 3 months sampled (Tukey F=4.89, DF=15, p<0.05) (Figure 2.3c). Similar results were found for the PV-tray communities (January 40+5species/tray). In addition, the species richness for the July PV-tray communities was significantly lower than that for the April PV-tray community (Tukey F=4.89, DF=15, p<0.05). Only in July was the total species richness for PL-tray communities (100+6species/tray) and the PV-tray communities (70+7species/tray) significantly different (Tukey F=4.89, DF=15, p<0.05). The July-only difference in total species richness between tray types is likely responsible for the significant interaction demonstrated in the two-factor ANOVA (Table 2.4). Species richness varied seasonally for 16 of the major groups (Table 2.3). Of these, 7 groups had significantly higher species richness in July than in January (see Figure 2.6a for example). Polychaeta, Asteroidea, Echinoidea and Hydrozoa had significantly higher species richness in April than in January and the Polychaeta also had higher species richness in April than in July and October (see Figure 2.6b for example). Five groups (Gastropoda, Holothuridea, Demospongiae, Turbellaria and Gymnolaemata) had the greatest species richness in the October communities relative to January or July (Turbellaria only) (see Figure 2.6c for example). In general, species richness was lowest in January for most of the major groups.  b Tray_Type PL PV  20  15  10  5  0 1 Jan  2 Apr  3 Jul  4 Oct  c 50  Gastropoda Spp. Richness ( # spp. / tray)  25  Polychaeta Spp. Richness ( # spp. / tray)  Malacostraca Spp. Richness ( # spp. / tray)  a  Tray_Type PL PV  40  30  20  10  0 1 Jan  2 Apr  3 Jul  4 Oct  15  Tray_Type PL PV  10  5  0 1 Jan  2 Apr  3 Jul  4 Oct  Figure 2.6. Seasonal mean species richness per fouling community (number species per tray) on suspended PL and PV trays for (a) Class Malacostraca, (b) Class Polychaeta and (c) Class Gastropoda.  29  A significant difference in species richness was detected between tray types for 4 major groups (Polychaeta, Bivalvia, Gastropoda and Foraminifera) indicating PL-trays had a greater number of species than PV-trays (see Figure 2.6b for example). In fact, for Polychaeta the July PL-tray communities had almost double the number of polychaete species as the July PV-tray communities. In addition to July, Polychaeta species richness in January was significantly higher in the PL-trays than in the PV-trays. These date-specific differences in species richness between the 2 tray types are reflected in the significant interaction results of the two-factor ANOVA for Polychaeta (Table 2.3). In summary, both total and major group species richness showed large seasonal variation and species richness varied between tray types for some major groups. The season with the greatest species richness was variable between the major groups. Six of the major groups (Polychaeta, Malacostraca, Bivalvia, Gastropoda, Gymnolaemata and Foraminifera) consistently had a large number of species present regardless of date or tray type. Class Polychaeta had the greatest number of species belonging to any major group, ranging from 11 species in the July PVtray samples to 37 species in April PL-tray samples. Of the 194 different species identified in the fouling communities, only 25 species were present on all sampling dates and in both tray types. Of these 25 species, 8 species were Malacostraca, 5 Polychaeta, 3 Bivalvia, 2 Gymnolaemata, 2 Gastropoda, 2 Foraminifera, 1 Maxillapoda, 1 Hydrozoa and 1 Ascidiacea.  2.3.1.4 Swartz Dominance Index (SDI) Mean SDI values for each fouling community are shown in Figure 2.7. Statistical analysis of the SDI values indicated that community dominance varied across season (ANOVA F(3, 15)=13.53, p<0.01), but there was no significant difference between tray types (ANOVA F(1, 15)=0.94, p<0.35). The greatest dominance (smallest SDI values) occurred in the July fouling communities (PL-tray 3.5+0.5SDI/tray and PV-tray 4.3+0.9SDI/tray) and the least dominance (largest SDI values) in the January and October communities for the PV-tray (11+1SDI/tray) and PL-tray (9SDI/tray) respectively (Figure 2.7) (Tukey F=4.94, p<0.05). The greater species dominance shown in the July tray fouling communities is supported by the abundance and biomass results (see sections 2.3.1.1 and 2.3.1.2), where there was an increase in the abundance and biomass of crustaceans, including Caprella mutica, Jassa staudei, Gammaridea sp., and Calanoid sp., and of the Bivalvia species Mytilus sp.  30  Dominance (SDI / tray)  15  Tray_Type PL PV  10  5  0 1 Jan n=3  2 Apr  3 Jul  n=3Date n=2  4 Oct n=3  Figure 2.7. Mean dominance per fouling community on suspended PL and PV trays (SDI per tray + SE).  2.3.1.5 Community analyses The results of the Bray-Curtis cluster analyses using species-abundance and species-biomass data support those of the univariate analyses: fouling communities do not differ between tray types, but they do vary between sampling dates. The cluster dendrogram in Figure 2.8 illustrates that species abundance in the samples collected from the 2 tray types on each date are more similar to each other than to the samples collected from either tray type on any other date. The cluster group including the July communities shows a dissimilarity of 0.95 to the cluster group including the January, April and October communities (linkage A). The null hypothesis that these 2 clusters were from the same community was rejected (p=0.010). In addition, the January communities were dissimilar to the October and April communities due to lower January abundances (Figure 2.8, linkage B, dissimilarity 0.89, p=0.006) and the October communities were dissimilar to the April communities (Figure 2.8, linkage C, dissimilarity 0.5 9, p=0.035). The results of the cluster analyses using species biomass data were similar to those using species abundance data, although the relative dissimilarity hierarchy of the dates was different. Based on species biomass data, the January tray communities had a dissimilarity of 0.99 to tray communities from other dates, although this result was only marginally significant (Figure 2.9, linkage A, p=0.055). This result is likely due to comparatively high within-group variability for biomass in both January tray communities (Figure 2.3b) combined with utilization of the SIGTREE analysis, which generated the null hypothesis probabilities by comparing within and  31  between group (community) variability (Nemec and Brinkhurst, 1988). The July tray communities had a 0.90 dissimilarity to October and April communities, due to the much higher biomass in the July communities (Figure 2.9, linkage B, p=0.005), and October communities had a 0.70 dissimilarity to April communities due to the higher biomass, distributed amongst fewer species in the October communities (Figure 2.9, linkage C, p<0.0001).  July-PL  F  July-PV January-PL  A  D  *  January-PV  B October-PL  G  *  C  October-PV  *  April-PL  E  April-PV  Dissimilarity  Figure 2.8. Hierarchical dendrogram showing the relative similarities of fouling communities using the Bray-Curtis dissimilarity index values of species abundance. Each letter (A through G) is associated with a cluster linkage. * represents a linkage where the non-parametric SIGTREE bootstrap method (200 simulations) detected the paired groups or “clusters” were significantly different from each other (Nemec and Brinkhurst, 1988).  January-PL  D  January-PV  A  F  July-PL July-PV  B  G  *  October-PL October-PV  C  *  E  April-PL April-PV  Dissimilarity  Figure 2.9. Hierarchical dendrogram showing the relative similarities of fouling communities using the Bray-Curtis dissimilarity index based on species biomass. Each letter (A through G) is associated with a cluster linkage. * represents a linkage where the non-parametric SIGTREE bootstrap method (200 simulations) detected the paired groups were significantly different from each other (Nemec and Brinkhurst, 1988).  32  2.3.2  Seasonal and spatial variation in water column parameters  Six water column variables (temperature, pH, salinity, dissolved oxygen, chlorophyll and turbidity) were measured from the surface to maximal depth (14.7m to 17.3m at farm stations, 23.3m at reference station) at all stations and on all dates. At all 5 stations, the July water temperatures were the highest recorded (14.3 to 15.7°C), followed by water temperatures in October (10.4°C), April (8.7 to 9.0°C) and January (7.9°C all stations) (for example, see Figure 2.10a). At all stations, water temperatures in January, April and October varied only slightly with depth. In July, a well-defined thermocline was present between 7.2m and 9.4m (temperature decreased 1.5°C) at the raft centre (C) station only (Figure 2.10a). Water column pH was slightly alkaline for all dates and all stations, with a range of 7.40 to 8.24. Spatial (between stations) variation in pH was minimal. There was very little variation in pH with increasing depth at any of the stations in January, April and October. In July, pH decreased slightly with depth at all stations with maximal decrease in pH at the reference station (8.20 to 7.80). At the C station, the rapid decrease in pH (8.07 to 7.95) occurred at the same depths (7.2 to 9.4m) as the thermocline (Figure 2.10b). Salinity water column profiles demonstrated little spatial variation between stations. Salinity was highest in April and lowest in July, with January and October salinities falling in between. January, April and October salinities varied only slightly with depth. July surface-water salinities, at all stations, were the lowest recorded in this study (24.8 to 26.2ppt) but increased with increasing depth (25.6 to 28.0ppt). At the C station in July, there was a halocline (salinity increased from 26.1 to 27.1ppt) present from 7.2m to 9.4m depth (Figure 2.10c). Water column dissolved oxygen (DO) did not vary spatially in January or October, but varied between stations in April. In July, DO levels within the raft were similar but DO was higher at the R station and lower at the NWF station. Dissolved oxygen was lowest in October (5.3 to 7.5mg/L all stations), marginally higher in April (7.9 to 9.6mg/L all stations), with water column DO for January and July falling in between (see Figure 2.10d for example). In January, April and October, DO decreased moderately with increasing depth. In July, DO levels decreased from the surface to maximal depth (11.6 to 5.6mg/L). In January and October, chlorophyll was consistently near 0µg/L throughout the water column at all 5 stations (see Figure 2.10e for example). Spatial variation occurred in April and July; chlorophyll levels at the SER and the NWF stations were lower in April than in July while chlorophyll levels at the NWR, C and R stations were generally higher in April than in July. In  33  April and July, chlorophyll profiles indicated a similar trend at all farm/raft stations with highest chlorophyll levels at 5m, followed by levels at 3m and then 11m. In contrast, the reference station had highest chlorophyll levels at 3m with lower levels at 5m and then 11m depth. Turbidity demonstrated very little spatial variation between stations on any of the sampling dates. In general, the October TSS levels throughout the water column were the lowest of all sampling dates, followed next by April or July and the highest TSS levels occurred in January (see Figure 2.10f for example). At most stations, April TSS levels increased from 3m (2.1 to 3.7mg/L) to 5m (3.9 to 6.2mg/L all stations) and decreased from 5 to 11m (2.2 to 3.8mg/L). In July, TSS levels demonstrated no general trend with depth. a  b  c  0  0  0  5  5  5  10  10  10  15  15  15  20  20  20 7  8  9  10 11 12 13 Temperat ure (°C)  14  15  7.6  7.7  7.8  7.9  8.0  26  8.1  d  e 0  0  5  5  5  10  10  10  15  15  15  6  7  8  9  Dissolved Oxygen (mg/ L)  10  31  32  20  20  20  28 29 30 Salinit y (ppt)  f  0  5  27  pH  0  2  4  6  Chlorophyll (µg/ L)  8  0  1  2  3  4  5  6  7  TSS (mg/ L)  Figure 2.10. Water column profiles in January , April , July and October at the raft centre (C) station. (a) Temperature, (b) pH, (c) salinity and (d) dissolved oxygen profiles were created using Sonde data, (e) chlorophyll and (f) turbidity (TSS) profiles were created using lab-analyzed Niskin water samples. Chlorophyll values for January are hidden by the October symbols.  Chlorophyll is commonly used as an indicator of plankton levels in the water column. TSS, however, measures all suspended particulates including biodeposits (feces and pseudofeces), sloughed organisms, organic material, as well as plankton. These 2 parameters are most informative when used together. For example, while chlorophyll was low in the winter months, TSS was high, which may be due to increased suspended particulate material as a result of storms or runoff. At the C station in April, chlorophyll was low at 3m while TSS was high suggesting  34  either that the particulates in the water column under the raft were not chlorophyll or that the oysters and fouling organisms were depleting the plankton at 3m depth. In fact, high chlorophyll values at 3m at the R station in April confirm the presence of high levels of plankton outside the farm and therefore, support the latter explanation. 2.4  DISCUSSION  The artificial substrates provided by the suspended oyster trays used in this study supported diverse fouling communities of marine invertebrates which demonstrated significant seasonal variation in abundance and biomass. The growth and reproduction of marine fouling communities is known to be affected by seasonal changes in the environment, such as fluctuations in temperature and salinity (Osman, 1977; Brown and Swearingen, 1998). These seasonal changes in key variables may impact phytoplankton, a food source for many fouling organisms and the cultured oysters, as well as organismal reproduction and larval dispersion (changes in water density) (Trujillo and Thurman, 2008). In this study, total abundance and biomass of fouling organisms increased from April to July, with the high April chlorophyll levels suggesting an abundance of plankton. The lower chlorophyll and TSS values recorded in July suggest reduced plankton, as well as larvae, available for growth and recruitment of fouling communities and the October fouling communities supported less abundance and biomass than those in July. In January, the lowest temperatures and chlorophyll levels coincided with the low abundance and biomass of fouling communities, suggesting little or no recruitment, reduced growth and organismal die off in the winter (Arakawa, 1990). Although the chlorophyll values reported were collected at a single point in time, date collected from the Strait of Georgia display similar seasonal patterns with high chlorophyll in the spring, particularly April, moderate values through the summer and fall, often with some peak blooms in the fall, and consistently low values in January (Masson and Pena, 2009). Seasonally high (summer) and low (winter) larval recruitment were reported for a fouling community on Perspex panels deployed for 2 years in Scotland (Turner and Todd, 1993). In Baja California, decreases in fouling coverage on suspended oyster shells deployed during the winter months (December to April) were also reported (Rodriguez and Ibarra-Obando, 2008). The seasonal changes in total abundance were influenced by changes in Bivalvia, which were dominated by Mytilus sp. and Hiatella arctica in July communities in particular. Mytilus edulis and H. arctica have also been reported as major contributors to community biomass on pearl nets containing scallops (Placopecten magellanicus) deployed from June to October in  35  Quebec (Claereboudt et al., 1994). Mytilus species have been reported as common dominants in fouling communities, but their total abundance is known to vary based on seasonal and annual recruitment in California and East Kamchatka (Scheer, 1945; Oshurkov, 1992). The abundance and biomass of the major taxonomic groups in the fouling communities showed seasonal trends, with greater values in the July communities and lower values in the January communities. Exceptions to these trends included Class Demospongiae (specifically Halichondria bowerbanki and Leucosolenia nautilia), for which abundance and biomass were greatest in the October fouling communities. H. bowerbanki has been reported as absent until the fall season in a study using ceramic plates in Beaufort, North Carolina (Sutherland and Karlson, 1977). The greater abundance and biomass of H. bowerbanki on oyster trays in October may be because only post-larval individuals that reproduce from July to mid-October were present on the trays rather than post-dormant individuals, as suggested by Fell and Jacob (1979) on eelgrass in the Mystic Estuary, Connecticut. Total species richness was lower in the winter than in the spring, summer or fall. In a study of fouling communities on Perspex panels in Scotland, Turner and Todd (1993) similarly reported fewer fouling species in winter than in summer. More species preferred to occupy PLtrays than PV-trays in July, specifically Polychaeta, Malacostraca and Foraminifera. In the literature, the polychaete Family Spirorbidae has been shown to have particular substrate preferences, although there was no comparison between plastic and polyvinyl. An Australian study showed this family preferred porous concrete and plywood to non-porous aluminium and fibreglass (Anderson and Underwood, 1994). In this study, Circeis armoricana was a Spirorbidae species present only on PL-trays in January and July. There is other literature indicating some fouling organisms preferentially choose particular substrates over others (e.g. Callow and Fletcher, 1994; Tyrrell and Byers, 2007; Erbland and Ozbay, 2008), however, it is unclear why particular species in this study preferentially choose PL-trays. PL-trays may be favoured over PV-trays by some species because PL-trays offer flat surfaces, while PV-tray surfaces are rounded. In this study, the total number of species identified within all the tray communities (194 species) exceeds the total number of species documented on artificial substrates in any of the aforementioned studies. Previous studies on fouling have reported total number of species ranging from 18 to 142 species on particular substrates, including pearl nets, oyster trays, slate, shell, wood and rubber. This noted difference in total species richness between studies is not  36  unexpected given that species richness may be dependant on substrate size (Osman, 1977) such that larger substrates (e.g. oyster trays) would attract more species than smaller settling plates. In addition, deployment date (season) and immersion time of the substrate have also been shown to be important in determining the number of species present (Anderson and Underwood, 1994). Species dominance within the fouling communities of both tray types was higher in July than on other sampling dates. This dominance was due to the seasonal high abundance of several Malacostraca species (e.g. Caprella mutica, Jassa staudei, and a Gammaridea sp.) and the Bivalvia species Mytilus sp. and H. arctica. The high abundance of C. mutica in July, in comparison with other sampling dates, may be related to this caprellid’s reproductive cycle. In this study, sea water temperatures (1m depth at the raft centre station) reached 13°C as early as May 24, and female C. mutica are known to brood at water temperatures between 13°C and 14°C and to have an average of 2 broods (1 released every 20 days) with up to 82 individuals per brood (Cook et al., 2007). Hence, it is likely that at least 47 days of suitable water temperatures for C. mutica reproduction occurred prior to the July sampling trip. July abundance of caprellids was 10 to 40 times more than that in April, which suggests that as many as 50% of the individuals in April were brooding females. As discussed earlier, the seasonal dominance of Mytilus sp. in the tray fouling communities of this study has also been reported in a number of other studies on temperate fouling communities (Scheer, 1945; Okamura, 1986; Hirata, 1987; Claereboudt et al., 1994). Biological interactions, such as predation and competition, may play an important role in determining the species composition of the fouling community (Mook, 1981; Richmond and Seed, 1991; Daka and Abby-Kalio, 2002). Predation has been shown to impact community structure (Dayton, 1977), largely because reduction of competitively dominant species through predation creates open spaces that are available for less competitive species to occupy (Osman, 1977). For example, both Okamura (1986) and Smedes and Hurd (1981) recorded the elimination of dominant mussels from fouling panels through predation by perch (Embiotica lateralis and Damalichthys vacca) and toad fish (Opsanus tau) respectively. In this study, perch (Brachyistus frenatus) were observed around the suspended oyster trays and were present in a single PL-tray in July and October. The presence of perch in and around the oyster trays suggests they are utilizing the seasonal abundance of fouling organisms, such as Mytilus sp. in July, as a food source. A number of other potential predators were present in the tray fouling communities (e.g. Pisaster ochraceus and Solaster stimpsoni).  37  The presence of fouling invertebrate groups such as the Ascidiacea, Polychaeta, Bivalvia, Demospongiae and Malacostraca can impact cultured oysters directly by colonizing the surface of the oysters or indirectly by colonizing the culture tray (Arakawa, 1990, Lesser et al., 1992). Species that colonize trays in encrusting forms and/or as colonies (e.g. sponge, colonial ascidians, encrusting bryozoans), as “block” species (species that are dense and can become large) (e.g. barnacles and mussels) or in extremely high abundance (solitary ascidians and some crustaceans) can disrupt the flow of water through the trays by partially or completely covering the tray mesh holes (Claereboudt et al., 1994; Lodeiros & Himmelman, 2000; Daka and Abby-Kalio, 2002). Alteration of the flow of water through the tray can impact the oysters’ access to oxygenated water and food. Ascidians, some polychaetes, barnacles, mussels and a variety of other filterfeeding organisms may compete directly with the cultured oysters by filtering similar-sized (2µm to 8µm) food particles (Buss and Jackson, 1981; Lesser et al., 1992; Ward et al., 1998; Ropert and Goulletquer, 2000). In this study, the Class Polychaeta (62 species) was undoubtedly one of the most influential major groups within the fouling communities, with some species having a potentially detrimental effect on the oysters. Filter-feeding and suspension-feeding polychaetes may compete for food (plankton) with the cultured oyster and in the tray fouling communities, members of 4 such polychaete families were present: Serpulidae (3 species), Sabellidae (6 species), Sabellariidae (1 species) and Spirorbidae (1 species). Of these 11 competitors only 2 species were sufficiently abundant to potentially compete with the cultured oysters for food. The serpulid, Pseudochitonopoma occidentalis was present in the communities on both tray types throughout the year, with peak abundance in July and low abundance in January. The sabellariid, Sabellaria cementarium, was also present on both tray types but was most abundant in the October fouling communities, although overall abundance was much lower than P. occidentalis. Given the low levels of chlorophyll and the peak abundance of most of the polychaete species in July, food competition between the oysters and polychaetes is possible if particles of comparable size are utilized. A study by Ropert and Goulletquer (2000) found that the Terebellidae polychaete Lanice conchilega could modify its feeding strategy from a deposit feeder to a deposit/suspension feeder and retain particles similar in size range to that of C. gigas. However, when feeding activity was standardized for weight, the activity of the polychaete was only 14% that of the oyster. There were 6 Terebellidae species identified in our study (predominantly in the April fouling  38  communities) but it is unknown whether they can retain particles in the same range as C. gigas. Ultimately it is unlikely that the Terebellidae species in this study substantially limited food availability to the oysters, as the April chlorophyll levels suggest plentiful food sources and these 6 species were present in relatively low abundance. Mussels (Mytilus) are filter feeders with a high filtration rate for a range of particle types and sizes that include those utilized by oysters and, therefore, have the potential to compete with oysters for food (Lesser et al., 1992). Mytilus may also disrupt water flow through the trays, reducing availability of food and oxygen to the oysters. In this study, Mytilus sp. was present largely as juveniles, in all fouling communities in relatively low abundance in January, April and October and in extremely high abundance in July (tens of thousands per tray). The great abundance of mussels present on the oyster trays undoubtedly had an effect on the oysters’ accessibility to resources, regardless of the size of mussels’. The removal of Mytilus before substantial growth is critical to oyster farm productivity. Ascidians, solitary and colonial, are filter feeders and when present in high abundance on oyster trays, can potentially compete with oysters for food. Studies on solitary ascidians have shown them to be active filter feeders (Fiala-Medioni, 1978; Young, 1989), filtering up to 444 litres/day (Young, 1989) and retaining particles >5µm (Randlov and Riisgard, 1979). Ascidians can compete with C. gigas and other filter-feeders not only because of their extremely high filtration rate but also because of their non-selective method of feeding (Rodriguez and IbarraObando, 2008). Because of the non-selective feeding method used by ascidians, they may consume large numbers of invertebrate larvae while actively filtering and, thus, may significantly affect fouling community structure (Cowden et al., 1984). Ascidians may also disrupt water flow through oyster cages at high abundance. The adverse effects of the solitary ascidian, Ciona intestinalis, on cultured oysters in Atlantic Canada has been reported by Carver et al. (2003). In this study, the solitary ascidian Corella inflata was present in all fouling communities and with high abundance in the July communities. Three non-native colonial ascidians, Botrylloides violaceus (July and October both tray types), Botryllus schlosseri (October PL-tray) and Didemnum sp. (July PL-tray), were also present in the tray fouling communities at very low abundances. In July, it is possible that the presence of C. inflata in the tray fouling communities had a significant negative effect on the water and food available to oysters. Fouling organisms, particularly non-native invertebrate species, have become increasingly important to both aquaculture operators concerned about damage to their product and ecologists  39  concerned about protecting local marine biodiversity and ecosystem function (Anil, 2006; Thébault and Loreau, 2006). In BC, to date, shellfish farmers have seen few of the potentially detrimental effects of invasive, non-native invertebrate species on cultured bivalves, although the presence of non-native species on aquaculture sites is increasing and their abundance is evident (pers. obs.). There were at least 11 non-native invertebrate species identified in the tray fouling communities in this study, 3 Malacostraca species (Caprella mutica, Jassa marmorata and Munna ubiquita), 3 Bryozoa species (Membranipora membranacea, Schizoporella unicornis and Bowerbankia gracilis), 1 Bivalvia species (Venerupis phillippinarum), 1 Turbellaria species (Pseudostylochus ostreophagus) and 3 Ascidiacea species (Botrylloides violaceus, Botryllus schlosseri and Didemnum sp.). The only non-native species with particularly high abundance was C. mutica (tens of thousands per tray), which had abundances second only to Mytilus sp. in the July fouling communities. These 11 invasive species are currently of minor concern to the oyster farm in this study, in regards to impacts on oyster health but these invasive species may collectively be altering the structure of the fouling communities by out-competing native fouling species that occupy similar niches (Cohen and Carlton, 1998; Stachowicz et al., 2002; Anil, 2006; Altman and Whitlatch, 2007) and their presence is cause for concern.  2.5  CONCLUSIONS  This comprehensive study provides a unique examination of the seasonal structure of fouling communities on oyster culture trays and contributes to our understanding of the role of deepwater, suspended oyster tray culture in supporting fouling communities. Fouling community composition is highly dependent on the time of year that the substrates are immersed and, based on the high abundance and biomass of fouling organisms recorded in July, growers with limited resources may want to focus on cleaning trays more frequently during this period of substantial settlement and growth. While plastic and polyvinyl substrates support similar fouling communities in terms of abundance and biomass, the fouling communities in the PL-trays had higher species richness than the PV-trays, such that species potentially harmful to oysters, including Botryllus schlosseri and Didemnum sp. were present on the PL-trays but absent on the PV-trays. Hence, although the PV-tray did not reduce fouling biomass, as previously suggested for this tray type, the absence of at least some of the invertebrate species potentially harmful to oysters (including some invasive species) from the PV-tray fouling communities provides evidence that this tray type may have other advantages for the grower. Regardless, the  40  development of an oyster tray that reduces the abundance and biomass of fouling on the trays, without compromising the density, health or growth rates of the oysters, should remain a focus for the oyster aquaculture industry. The regional predictability of fouling communities is of ongoing relevance to both oyster farmers and ecologists. Unfortunately, a number of factors including spatial and temporal changes in coastal conditions and thus larval recruitment, the introduction of non-native species, and irregular or unpredictable natural disturbances can make it difficult to predict the short-term and long-term development of fouling communities. Future studies should focus on annual and spatial fluctuations in local fouling communities (e.g. farms in close proximity), close examination of larval settlement (e.g. using fouling panels) and a farm-based study, such as the one here using oyster trays, to better understand the seasonal and spatial distribution of fouling organisms, the local availability of larvae for settlement, larval substrate preference or avoidance, and the significant effects of fouling on the growth and survival of cultured oysters.  41  LITERATURE CITED Aleem, A.A., 1957. Succession of marine fouling organisms on test panels immersed in deepwater at la Jolla, California. Hydrobiologia 11: 40-58. Altman, S., Whitlatch, R.B., 2007. Effects of small-scale disturbance on invasion success in marine communities. Journal of Experimental Marine Biology and Ecology 342: 15-29. Anderson, M.J., Underwood, A.J., 1994. Effects of substratum on the recruitment and development of an intertidal estuarine fouling assemblage. Journal of Experimental Marine Biology and Ecology 184: 217-236. Anil, A.C., 2006. A perspective of marine bioinvasion. In multiple dimensions of global environmental change, TERI Press, India. Arakawa, K.Y., 1990. Competitors and fouling organisms in the hanging culture of the Pacific oyster, Crassostrea gigas (Thundberg). Marine and Freshwater Behaviour and Physiology 17: 67-94. Arar, J.E., 1997. Method 446.0 - In vitro determination of chlorophyll a, b, c1 + c2 and pheopigments in marine and freshwater algae by visible spectrophotometry. National Exposure Research Laboratory, U.S. Environmental Protection Agency, 12p. Barbour, M.T., Gerritsen, J., 1998. Subsampling of benthic samples: A defense of the fixed-count method. Journal of the North American Benthological Society 15: 386-391. BCSGA, 2009. British Columbia Shellfish Growers Association: Industry Encyclopaedia. http://www.bcsga.ca/encyclopedia_index.php. Bram, J.B., Page, H.M., Dugan, J.E., 2005. Spatial and temporal variability in early successional patterns of an invertebrate assemblage at an offshore oil platform. Journal of Experimental Marine Biology and Ecology 317: 223-237. Bray, J.R., Curtis, J.T., 1957. An ordination of the upland forest communities of southwestern Wisconsin. Ecological Monographs 27: 325-349. Brown, K.M., Swearingen, D.C., 1998. Effects of seasonality, length of immersion, locality and predation on an intertidal fouling assemblage in the Northern Gulf of Mexico. Journal of Experimental Marine Biology and Ecology 225: 107-121. Buss, L.W., Jackson, J.B.C., 1981. Planktonic food availability and suspension-feeder abundance: Evidence of in situ depletion. Journal of Experimental Marine Biology and Ecology 49: 151-161. Callow, M.E., Fletcher, R.L., 1994. The influence of low surface energy materials on bioadhesion- A review. International Biodeterioration and Biodegradation 34: 333-348.  42  Carver, C.E., Chisholm, A., Mallet, A.L., 2003. Strategies to mitigate the impact of Ciona intestinalis (l.) biofouling on shellfish production. Journal of Shellfish Research 22: 621631. Chalmer, P.N., 1982. Settlement patterns of species in a marine fouling community and some mechanisms of succession. Journal of Experimental Marine Biology and Ecology 58: 7385. Claereboudt, M.R., Bureau, D., Côté, J., Himmelman, J.H., 1994. Fouling development and its effect on the growth of juvenile giant scallops (Placopecten magellanicus) in suspended culture. Aquaculture 121: 327-342. Cohen, A.N., Carlton, J.T., 1998. Accelerating invasion rate in a highly invaded estuary. Science 279: 555-558. Cook, E.J., Jahnke, M., Kerckhof, F., Minchin, D., Faasse, M., Boos, K., Ashton, G., 2007. European expansion of the introduced amphipod Caprella mutica Schurin 1935. Aquatic Invasions 2: 411-421. Cowden, C., Young, C.M., Chia, F.S., 1984. Differential predation on marine invertebrate larvae by two benthic predators. Marine Ecology Progress Series 14: 145-149. Daka, E.R., Abby-Kalio, N.J., 2002. The effects of caging on the colonization of fouling organisms in the upper Bonny estuary. Journal of Applied Sciences and Environmental Management 6: 29-33. Dayton, P.K., 1971. Competition, disturbance, and community organization: The provision and subsequent utilization of space in a rocky intertidal community. Ecological Monographs 41: 351–389. Dean, T.A., Hurd, L.E., 1980. Development in an estuarine fouling community: The influence of early colonists on later arrivals. Oecologia 46: 295-301. DFO, 2005. Juan de Fuca Strait to Strait of Georgia #3462. DFO, Canadian Hydrographic Service. DFO, 2006. Assessing Habitat Risks Associated with Bivalve Aquaculture in the Marine Environment. DFO, Canadian Science Advisory Secretariat, Science Advisory Report 2006/005. Erbland, P.J., Ozbay, G., 2008. A comparison of the macrofaunal communities inhabiting a Crassostrea virginica oyster reef and oyster aquaculture gear in Indian River Bay, Delaware. Journal of Shellfish Research 27: 757-768. Enright, C., 1993. Control of fouling in bivalve aquaculture. World Aquaculture 24(4): 44-46. Fiala-Medioni, A., 1978. Filter-feeding ethology of benthic invertebrates (Ascidians). IV. Pumping rate, filtration rate, filtration efficiency. Marine Biology 48: 243-249.  43  Fell, P.E., Jacob, W.F. 1979. Reproduction and development of Halichondria sp. in the Mystic Estuary, Connecticut. Biological Bulletin 156: 62-75. Fry, W.G., 1975. Raft fouling in the Menai Strait, 1963-1971. Hydrobiologia 47: 527-558. Guenther, J., Southgate, P.C., de Nys, R., 2006. The effect of shell size on accumulation of fouling organisms on the Akoya pearl oyster Pinctada fucata (Gould). Aquaculture 253: 366-373. Hirata, T., 1987. Succession of sessile organisms on experimental plates immersed in Nabeta Bay, Izu Pennisula, Japan. II. Succession of invertebrates. Marine Ecology Progress Series 38: 25-35. Hintze, J. 2007. NCSS and GESS. NCSS, LLC. Kaysville, Utah. www.ncss.com. Huguenin, J.E., Huguenin, S.S., 1982. Biofouling resistant shellfish trays. Journal of Shellfish Research 2: 41-46. Lesser, M.P., Shumway, S.E., Cucci, T., Smith, J., 1992. Impact of fouling organisms on mussel rope culture: Interspecific competition for food among suspension-feeding invertebrates. Journal of Experimental Marine Biology and Ecology 165: 91-102. Lodeiros, C.J.M., Himmelman, J.H., 2000. Identification of factors affecting growth and survival of the tropical scallop Euvola (Pecten) ziczac in the Golfo de Cariaco, Venezuela. Aquaculture 182: 91-114. Masson, D., Pena, A., 2009. Chlorophyll distribution in a temperate estuary: The Strait of Georgia and Juan de Fuca Strait. Estuarine, Coastal and Shelf Science 82: 19-28. Miron, G., Landry, T., MacNair, N., 2002. Predation potential by various epibenthic organisms on commercial bivalve species in Prince Edward Island: Preliminary results. Canadian Technical Report of Fisheries and Aquatic Sciences 2392: 1-44. Mook, D.H., 1981. Effects of disturbances and initial settlement on fouling community structure. Ecology 62: 522-526. Moran, S.B., Charette, M.A., Pike, S.M., Wicklund, C.A., 1999. Differences in seawater particulate organic carbon concentration in samples collected using small- and large-volume methods: the importance of DOC adsorption to the filter blank. Marine Chemistry, 67: 3342. Moura, A., Boaventura, D., Curdia, J., Carvalho, S., Cancela de Fonseca, L., Leitao, F.M., Santos, M.N., Monteiro, C.C., 2007. Effect of depth and reef structure on early macrobenthic communities of the Algrave artificial reefs (southern Portugal). Hydrobiologia 580: 173180.  44  Nemec, A.F.L., Brinkhurst, R.O., 1988. Using the bootstrap to assess statistical significance in the cluster analysis of species abundance data. Canadian Journal of Fisheries and Aquatic Science 45: 965-970. Okamura, B., 1986. Formation and disruption of aggregations of Mytilus edulis in the fouling community of San Francisco Bay, California. Marine Ecology Progress Series 38: 275-282. Oshurkov, V.V., 1992. Succession and climax in some fouling communities. Biofouling 6: 1-12. Osman, R.W., 1977. The establishment and development of a marine epifaunal community. Ecological Monographs 47: 37-63. Randlov, A., Riisgard, H.U., 1979. Efficiency of particle retention and filtration rate in four species of ascidians. Marine Ecology Progress Series 1: 55-59. Richmond, M.D., Seed, R., 1991. A review of marine macrofouling communities with special reference to animal fouling. Biofouling 3: 151-168. Rodriguez, L.F., Ibarra-Obando, S.E., 2008. Cover and colonization of commercial oyster (Crassostrea gigas) shells by fouling organisms in San Quintin, Mexico. Journal of Shellfish Research 27: 337-343. Ropert, M., Golletquer, P., 2000. Comparative physiological energetics of two suspension feeders: Polychaete annelid Lanice conchilega (Pallas 1766) and cupped oyster Crassostrea gigas (Thundberg 1795). Aquaculture 181: 171-189. Scheer, B.T., 1945. T he development of marine fouling communities. Biological Bulletin 89: 103-121. Smedes, G.W., Hurd, L.E., 1981. An empirical test of community stability: Resistance of a fouling community to a biological patch-forming disturbance. Ecology 62: 1561-1572. Sneath, P.H.A., Sokal, R.R., 1973. “Numerical taxonomy: The principles and practice of numerical classification”. W.H. Freeman, San Francisco, 573 pp. Stachowicz, J.J., Fried, H., Osman, R.W., Whitlatch, R.B., 2002. Biodiversity, invasion resistance and marine ecosystem function: Reconciling pattern and process. Ecology 83: 2575-2590. Sutherland, J.P., Karlson, R.H., 1977. Development and stability of the fouling community at Beaufort, North Carolina. Ecological Monographs 47: 425-446. Swartz, R.C., 1978. Techniques for sampling and analyzing the marine macrobenthos. U.S. Environmental Protection Agency (EPA), Doc. EPA-600/3-78-030, EPA, Corvallis Oregon. 27p. Trujillo, A.P., Thurman, H.V., 2008. Essentials of oceanography. 9th ed. Pearson Prentice Hall, New Jersey.  45  Turner S.J., Todd, C.D., 1993. The early development of epifaunal assemblages on artificial substrata at two intertidal sites on an exposed rocky shoe in St. Andrews Bay, N.E. Scotland. Journal of Experimental Marine Biology and Ecology 166: 251-272. Ward, J.E., Levinton, J.S., Shumway, S.E., Cucci, T., 1998. Particle sorting in bivalves: In vivo determination of the pallial organs of selection. Marine Biology 131: 283-292. Young, C.M., 1989. Larval depletion by ascidians has little effect on settlement of epifauna. Marine Biology 102:481-489.  46  CHAPTER THREE. Seasonal comparison of invertebrate fouling communities associated with Crassostrea gigas cultured in two different tray types 2 3.1  INTRODUCTION  Bivalve aquaculture sites provide an ideal environment for the settlement and growth of diverse fouling communities, consisting particularly of sessile invertebrates, through colonization of the farming equipment and the cultivated species (Wahl, 1989; DFO, 2006). In the marine environment, naturally occurring hard substrates, such as rock and shell, are limited (Lesser et al., 1992) resulting in sessile organisms that require suitable substrate often occupying artificial substrates (docks, ropes, etc.). Naturally occurring oyster reefs support some of the most diverse hard-substrate communities by providing complex three-dimensional habitat with an abundance of suitable substrate, including protective crevices between and among the oysters (McKindsey et al., 2006; Erbland and Ozbay, 2008). Oyster culture farms, particularly deep-water suspended operations, are thought to provide habitat similar to that of oyster reefs (Erbland and Ozbay, 2008), although oyster farms have an artificial substrate component which may affect the composition of organisms that colonize. The introduction of cultured bivalves into coastal ecosystems can potentially offer the complex substrate required to maintain or increase local marine invertebrate biodiversity (Gray, 1997). This potential benefit of bivalve aquaculture may be tempered, however, by the unintentional introduction of non-native species with aquaculture or by providing an ideal habitat for introduced species to persist (Stachowicz et al., 2002). The composition of invertebrate fouling communities associated with cultured bivalve species varies seasonally and is strongly influenced by larval recruitment, competition, predation, species life histories, and local environmental conditions (Lesser et al., 1992; Dalby and Young, 1993; Mazouni et al., 2001). Larval recruitment is dependant on the presence, dispersal and settlement of particular species of larvae, which in turn are strongly influenced by local water conditions and currents, competition and predation (Osman, 1977; Walters and Wethey, 1991; Woodin, 1991). Fouling is important to growers because it can potentially reduce bivalve growth and survival, alter shell morphology, affect overall bivalve appearance, and ultimately decrease the profit margin (Murad and Mohammad, 1976; Taylor et al., 1997; Carver et al., 2003; Lodeiros and Garcia, 2004; Guenther et al., 2006). There have been very few studies that have looked 2  A version of this chapter will be submitted for publication. Switzer, S.E., Barnes, P.A., McKinley, R.S. Seasonal comparison of invertebrate fouling communities associated with Crassostrea gigas cultured in two different tray types.  47  closely at fouling communities inhabiting cultured bivalve species, possibly due to the challenging nature of undertaking this type of research, particularly with respect to taxonomy. Fouling organisms that grow on cultured bivalves and which are considered a nuisance to growers include barnacles (Balanus spp.), ascidians, polychaetes (Serpulidae spp., Spionidae spp.) and mussels (Mytilus spp.) (Arakawa, 1990; Lesser et al., 1992; Mazouni et al., 2001; Carver et al., 2003). Several antifouling treatments are available for growers, including chemical (brine dip), mechanical (exposure to heat, sun, hot water or freshwater and manual removal) or biological (introduction of the natural predators of the fouling organisms) treatments. Treatment options can be costly to growers (increased labour costs) and may effect survival of the treated bivalves (Arakawa, 1990; Lodeiros and Garcia, 2004; Ross et al., 2004). Recently, metal trays with polyvinyl coating (PV) have started replacing the “traditional” plastic trays (PL) due to their increased oyster holding densities combined with the belief that they reduce fouling. In combination with investigating the efficacy of using alternative tray types, an increased understanding of the seasonal composition of fouling communities may allow growers to identify the optimal time of year to employ preventative measures, thereby increasing cost effectiveness. This study set out to examine the seasonal variation in the invertebrate fouling community occupying the surface of Pacific oysters (Crassostrea gigas), cultured in the PL and PV tray types, at a deep-water suspended oyster farm in British Columbia, Canada. The experimental trays were subject to standard handling procedures for the aquaculture site and the fouling communities on seeded oysters in the trays were examined after 3, 6 and 12 months. The results of this study will aid growers in determining whether tray type affects the fouling communities on the oysters and, if seasonal changes in the fouling community are observed, whether these changes can be utilized in targeting the most effective time of year to employ antifouling methods.  3.2  METHODS  3.2.1  Study site and experimental design  The study site was a deep-water suspended Pacific oyster (C. gigas) farm located between Saltspring Island and Vancouver Island in Sansum Narrows, BC (see Figure 2.1a). The oyster farm was positioned on the east side of the Narrows, close to Saltspring Island, in the south end of a small embayment (see Figure 2.1b). The standard experimental raft (Figure 2.1b inset) used in  48  this study was located in the southeastern corner of the farm, on the north side of a group of 3 standard rafts. Two experimental tray types, a “traditional” plastic tray (PL) and a modern metal tray with a polyvinyl coating (PV), were used in this study (see Section 2.2.1 and Figure 2.2 for tray details). The experimental trays were seeded at the same time as the farm trays and followed the farm’s standard seeding practice. In October 2006, trays were seeded at high density (120 oysters per tray) and after 3 months the density reduced to 60 oysters per tray. The oyster stacks were suspended from the experimental raft at a minimum depth of 1.5m and in January 2007, April 2007 and October 2007, the 6 experimental stacks were retrieved and the fouling collected from all of the oysters in the third tray in each experimental stack. Oysters in the experimental trays were tumbled following removal of fouling. Oyster tumbling, in which oysters are processed through a mechanical tumbler, forcefully removes excess shell and allows the oyster to focus on meat growth. The procedure at this farm was to tumble the oysters at both 3 months (January 2007) and 6 months (April 2007) after seeding. On each sampling trip, 5 oysters from each experimental tray were removed and measured to determine growth and changes in surface area. In addition, water column data was collected on each sampling trip.  3.2.2  Field methods  In January, April, July and October 2007, fouling was collected from oysters in the 6 experimental trays by washing the surface of each oyster with a wash bottle and then removing any remaining organisms using forceps and a scalpel. Fouling was placed into seawater-filled totes that were fitted with a spout and a 500µm Nitex filter to allow water to be replaced without the loss of organisms. At each collection date, after fouling was removed, the oysters in each experimental tray were counted and 5 oysters were randomly chosen for determining average oyster length, width, height and weight. These 5 oysters were not returned to their experimental tray, but were transported to the laboratory for measurement and then were frozen for later estimation of surface area. Water column parameters (temperature, pH, chlorophyll, turbidity, salinity and dissolved oxygen) were monitored as described in Section 2.2.2.  49  3.2.3  Laboratory methods  In the laboratory, located at the Centre for Shellfish Research (Vancouver Island University, Nanaimo, BC), fouling samples were filtered using a 500µm Nitex screen, preserved in 10% buffered formalin for 3 days and then transferred to 75% ethanol for storage. Preserved fouling samples were processed by removing all organisms from the organic debris and shell using a dissecting microscope, identifying all organisms to species level, counting the number of individuals for each species and then placing them within a size category based on the known adult size of that species. Representatives of each size category for each species were weighed (wet weights) to estimate biomass. Colonial species counts were estimated by determining the number and size of fragments present and then extrapolating to determine the possible number of colonies. January, April and October 2007 fouling samples were processed by removing the large organisms, identifying them and then splitting the remaining material into 2 equal portions (Barbour and Gerritsen, 1998). To ensure that our sampling procedure resulted in 2 equal representative samples 4 of the January samples were split and both portions immediately processed. Our findings indicated there were no significant differences between the 2 subsamples arising from the same original sample. All fractioned fouling data were multiplied, or the fractions were added together if both halves were processed, to determine totals for the whole samples. Surface area was estimated for 30 frozen oysters selected from the oysters collected on each sampling trip from each experimental tray. The oysters used for these estimates were selected to provide a good representation of the range of oyster surface areas on all dates and were of variable sizes and shapes. Surface area was measured by covering both sides of each oyster with tinfoil, ensuring that the foil was fitted as closely as possible into each crevice, trimming the foil along the outer edges of the shells and then removing the foil and flattening it onto grid paper (Murad and Mohammad, 1976). The total area of foil was equivalent to the surface area of the oyster (m2). Water column data were considered using methods outlined in Section 2.2.3.  3.2.4  Data analysis  On each sampling date, each fouling community was represented by 3 replicate samples. Because the total surface area of oysters (i.e. available substrate for fouling organisms) varied between trays and increased with oyster growth throughout the study, the oyster fouling data was  50  standardized using oyster surface area (m2). The relationship between oyster surface area and individual oyster growth parameters (length, width, height and weight) was explored using linear regression. The growth parameter that best explained increases in oyster surface area, length (R2=0.8048), was then used to estimate the total surface area of the oysters within each experimental tray using the mean oyster length and the total number of oysters in the tray. Oyster fouling abundance, biomass and species richness values were then standardized using total oyster surface area estimates, which were calculated using the defined linear relationship between surface area (y) and oyster length (x), where y = (-0.0024) + (0.1252) * x. Fouling community composition on oysters grown in the 2 tray types was evaluated using measures of abundance (total number of individuals per m2, number of individuals per major taxonomic group per m2 and number of individuals per species per m2), biomass (total organismal weight per m2, organismal weight per major taxonomic group per m2 and organismal weight per species per m2) and species richness (total number of species per m2, and per major taxonomic group per m2). For the purpose of data analysis, fouling community organisms were separated into major taxonomic groups, which included grouping invertebrates into taxonomic classes and non-invertebrates into common groups (e.g. Foraminifera). The large number of species identified required that we use major taxonomic groups rather than species for most analyses. Community dominance was measured for each tray of oysters using the Swart Dominance Index (SDI), a measure of the minimum number of species that account for 75% of the total abundance (Swartz, 1978). Abundance, biomass and species richness (total and for each major taxonomic group as described above), as well as SDI, were compared between dates and tray types using a TwoFactor Analysis of Variance (ANOVA α=0.050), followed by post-hoc multiple comparison Tukey Tests (Tukey α=0.050). Abundance and biomass data were log-transformed to satisfy the assumption of normality for the parametric ANOVA and Tukey tests. Using NCSS statistical software (Hintze, 2007), the parametric normality tests used were Shapiro-Wilk, AndersonDarling, D’Agostino Skewness and D’Agostino Kurtosis. In some cases, results of the ANOVA (pooled data) conflicted with those of the Tukey post-hoc tests (non-pooled data). Because the ANOVA uses pooled data, the degrees of freedom are higher than those of the Tukey tests, increasing the power of the test and allowing smaller differences to be detected. Such conflict between the results of the 2 tests also arises when the dataset contains many zeroes (i.e. species or major groups are absent) and/or when within-community variation is high. Under these  51  conditions, the power of the Tukey tests is reduced and it cannot detect differences between groups due to the high variability around the mean. In this study, the results of the ANOVA were always reported and when the Tukey tests could help explain the ANOVA significant differences these results were also reported. Multivariate analyses, using species abundance and species biomass data, were used to compare all fouling communities. Bray-Curtis dissimilarity coefficients (Bray and Curtis, 1957) were calculated separately for species abundance and species biomass data for all possible pairs of the 6 fouling communities. The dissimilarity coefficients were arranged graphically by creating a hierarchical cluster dendrogram using an unweighted pair-group mean average sort algorithm (Sneath and Sokal, 1973). For each linkage in the dendrogram the between-group versus the within-group variability was analyzed using a non-parametric bootstrap method using 200 simulations for each analysis (SIGTREE - Nemec and Brinkhurst, 1988). This method tests the null hypothesis for each linkage independently: that the 2 clusters (communities) being compared are the same community (α=0.050)  3.3  RESULTS  3.3.1  Composition of fouling communities on tray-cultured oysters  Oyster fouling abundance, biomass and species richness values were standardized per tray using estimates of the surface area of all oysters in the tray (see Methods), with parameters reported as per m2. Total surface area of the oysters was greatest in January trays because experimental trays were initially seeded with 120 oysters in October but densities were reduced to 60 oysters after sampling in January and 5 oysters were removed each sampling trip thereafter. During the April to October period, total surface area of the oysters increased slightly, due to oyster growth (Figure 3.1). Organisms belonging to 21 major taxonomic groups were identified in the oyster fouling communities sampled, from both tray types, in January, April and October: 20 invertebrate classes as well as the non-invertebrate group Foraminifera (Table 3.1). All fouling communities combined supported a total of 133,792 individuals belonging to 172 invertebrate species and 10 Foraminifera species with a total biomass of 312g. The fouling communities from oysters in each tray type, for each of the 3 sampling dates, were compared using univariate analyses (ANOVA and Tukey’s post-hoc tests) of abundance (log transformed), biomass (log transformed), species richness and a measure of dominance (Swartz Dominance Index), as well as multivariate analysis  52  (hierarchical cluster analysis using Bray-Curtis dissimilarity coefficients) of species abundance and species biomass.  Total Oyster Surface Area (sq.m)  1.0 Tray_Type PL PV  0.8  0.6  0.4  0.2  0.0 1 January n=3  2 April  3 October  n=3 Date  n=3  Figure 3.1. Mean total oyster surface area per fouling community on oysters suspended in PL and PV-trays (m2 + SE).  3.3.1.1 Abundance Total abundance of individuals differed between sampling dates but not between tray types (Table 3.2). Specifically, October fouling communities for both tray types had significantly higher abundance than either January or April communities (Tukey F=4.75, DF=12, p<0.05) while January and April communities had very similar total abundance of individuals (Figure 3.2a). Similar to total abundance, the abundance of each major taxonomic group varied between sampling dates for 19 of the 21 major taxonomic groups (Table 3.1). For nearly all of the 19 major groups, the abundance of the October oyster fouling communities (1 or both tray types) was greater than that of the January and April communities. The Class Polychaeta was the most abundant group, in both tray types, in January, April and October with Platynereis bicanaliculata, being the most abundant species (Figure 3.3a). Class Malacostraca had a relatively low abundance in January but a large increase in abundance in the April and October communities (Figure 3.3b); the increase was strongly influenced by the amphipod Aorides intermedius. Class Ostracoda, Ophiuroidea and Demospongiae showed a difference in abundance between tray types. Demospongiae abundance in the October PL-trays was higher than in the October PV-trays (Tukey F=4.75, DF=12, p<0.05) (Figure 3.3c) as a result of Halichondria bowerbanki. ANOVA results also indicated that the interaction between date and tray type influenced the abundance of  53  Class Ophiuroidea and Demospongiae, as well as Class Holothuridea which was marginally significant (Table 3.1). Table 3.1. Summary of two-factor ANOVA results (p-values) for the 21 major groups for abundance (log (x+0.1) transformed) (number individuals per m2), biomass (log (x+0.1) transformed) (g per m2) and species richness (number species per m2) for 2 factors (date and tray type) and the interaction between the 2 factors. * indicates a significant pvalue and ** indicates ANOVA results that were not supported by Tukey post-hoc results. Abundance  Biomass  Species Richness  Major Group  Date  Tray Type  Interaction  Date  Tray Type  Interaction  Date  Tray Type  Interaction  Polychaeta  <0.01 *  0.58  0.61  <0.01 **  0.93  0.93  <0.01 *  0.88  0.30  Malacostraca  <0.01 *  0.85  0.71  <0.01 *  0.73  0.62  <0.01 *  0.55  0.79  Maxillapoda  <0.01 *  0.29  0.58  <0.01 *  0.39  0.36  <0.01 *  0.72  0.34  Ostracoda  <0.01 *  0.02 **  0.16  0.01 **  0.25  0.30  <0.01 *  0.74  0.59  Pycnogonida  <0.03 **  0.46  0.58  0.20  0.29  0.33  <0.05 **  0.69  0.85  Bivalvia  <0.01 *  0.28  0.98  <0.01 *  0.77  0.99  <0.04 **  0.93  0.84  Gastropoda  <0.01 *  0.09  0.50  <0.01 *  0.19  0.26  <0.01 *  0.57  0.98  Polyplacophora  0.31  0.65  0.94  0.06  0.13  0.30  0.24  0.99  0.94  Asteroidea  <0.01 *  0.34  0.65  <0.01 *  0.80  0.96  0.27  0.33  0.91  Echinoidea  <0.01 **  0.29  0.63  0.47  0.32  0.41  <0.01 *  0.58  0.31  Holothuridea  <0.01 *  0.98  0.07  <0.01 *  0.02 *  0.01 *  0.07  0.27  0.16  Ophiuroidea  <0.01 *  0.05 **  0.03 *  0.39  0.34  0.40  <0.01 *  0.09  0.07  Ascidiacea  <0.01 *  0.94  0.41  <0.01 *  0.22  0.30  0.09  0.47  0.11  Demospongiae  <0.01 *  <0.01 *  <0.01 *  <0.01 *  <0.01 *  <0.01 *  <0.01 *  0.53  0.12  Gymnolaemata  <0.01 *  0.60  0.48  <0.01 *  0.01 **  0.12  <0.01 *  0.04**  0.36  Anopla  <0.01 *  0.51  0.64  <0.01 **  0.71  0.86  <0.01 *  0.16  0.14  Enopla  <0.01 *  0.17  0.09  0.06  0.69  0.40  <0.01 **  0.34  0.19  Anthozoa  0.11  0.67  0.12  0.80  0.24  0.35  0.02**  0.27  0.92  Hydrozoa  <0.01 *  0.49  0.09  0.33  0.04 **  0.76  <0.01 *  0.84  0.27  Turbellaria  <0.01 *  0.89  0.49  0.29  0.60  0.52  <0.01 *  0.41  0.04*  Foraminifera  <0.01 *  0.77  0.92  <0.01 **  0.83  0.86  <0.01 **  0.64  0.74  Table 3.2. Summary of two-factor ANOVA results for abundance (log (x+0.1) transformed) (total number individuals per m2), biomass (log (x+0.1) transformed) (total g per m2) and species richness (total number species per m2) for 2 factors (date and tray type) and for the interaction between the 2 factors. * indicates a significant p-value Abundance  F-Ratio p-value Power (α=0.05)  Biomass  Date  Tray Type  Interaction  Species Richness  Date  Tray Type  Interaction  Date  Tray Type  Interaction  56.74  1.84  0.00  53.95  1.66  2.14  15.46  0.35  0.29  <0.01 *  0.20  0.99  <0.01 *  0.22  0.16  <0.01 *  0.57  0.75  1.00  0.24  0.05  1.00  0.22  0.35  0.99  0.08  0.86  54  a  a  c 3.0  Tray_Type PL PV  4.0  3.0  2.0  1.0  Tray_Type PL PV  2.0  1.0  0.0  0.0 1 January n=3  1 January n=3  2 3 April October Date n=3 n=3  250  Total Species Richness (# spp. / sq.m)  log_Total Abundance (# indiv / sq.m)  5.0  b  log_Total Biomass (g / sq.m)  a  Tray_Type PL PV  200  150  100  50  0  2 October 3 April Date n=3 n=3  1 January n=3  2 April  Date n=3  3 October n=3  Figure 3.2. Seasonal changes in fouling communities on oysters suspended in PL and PV-trays. (a) Mean total abundance (log transformed) per fouling community (number of individuals per m2 + SE), (b) Mean total biomass (log transformed) per fouling community (g per m2 + SE) and (c) Mean total species richness per fouling community (number of species per m2 + SE).  b 5  4  Tray_Type PL PV  3  2  1  0 1  Jan  2  Apr Date  3  Oct  5  4  c Tray_Type PL PV  3  2  1  0 1  Jan  2  Apr Date  3  Oct  log_Demospongiae Abund. (# indiv / sq.m)  a log_Malacostraca Abundance (# indiv / sq.m)  log_Polychaeta Abundance (# indiv / sq.m)  a  5  4  Tray_Type PL PV  3  2  1  0 1  Jan  2  Apr Date  3  Oct  Figure 3.3. Seasonal mean abundance (log transformed) per fouling community (number individuals per m2) on oysters suspended in PL and PV-trays for (a) Class Polychaeta, (b) Class Malacostraca and (c) Class Demospongiae.  3.3.1.2 Biomass Total biomass differed between sampling dates, but not between tray types (Table 3.2). Similar to total abundance results (see Figure 3.2a), the October communities had greater biomass (both tray types), than the January and April communities (Tukey F=4.75, DF=12, p<0.05), while the January and April communities were similar to each other (Figure 3.2b).  55  Statistical results indicated that the biomass of 13 major groups were significantly different between sampling dates (Table 3.1). The dominant seasonal trends in major group biomass were low biomass in January communities and a much higher biomass in October communities, with April biomass falling between those of January and October. Some fouling community major groups (Malacostraca, Ascidiacea, Demospongiae (Figure 3.4a), Gymnolaemata (Figure 3.4b) and Enopla) also had significantly different biomass between January and April for one or both tray types. b 2  Tray_Type PL PV 1  -1  -2 1 Jan  2 Apr  Date  3 Oct  c 2  2  Tray_Type PL PV  log_Holothuridea Biomass (g / sq.m)  a log_Gymnolaemata Biomass (g / sq.m)  log_Demospongiae Biomass (g / sq.m)  a  1  0  -1  -2  Tray_Type PL PV  1  -1  -2 1  Jan  2  Apr  Date  3  Oct  1  Jan  2  Apr Date  3  Oct  Figure 3.4. Seasonal mean biomass (log transformation) per fouling community (g per m2) on oysters suspended in PL and PV-trays for (a) Class Demospongiae, (b) Class Gymnolaemata and (c) Class Holothuridea.  Differences in the biomass of 4 major groups were found between the 2 tray types: Holothuridea (Figure 3.4c), Demospongiae (Figure 3.4a), Gymnolaemata (Figure 3.4b) and Hydrozoa (Table 3.1). These groups had greater biomass in the October PL-tray communities than in the October PV-tray communities (Tukey F=4.75, DF=12, p<0.05). Interestingly, the Demospongiae biomass was almost identical in the 2 trays in April, but in October, biomass in the PV-tray was lower than in April while the PL-tray biomass was higher. The increase (PL-trays) and decrease (PV-trays) in Demospongiae biomass between the April and October sampling dates was mirrored in the abundance results (see section 3.3.1.1) suggesting seasonal growth of the 2 species of sponge, Halichondria bowerbanki and Leucosolenia nautilia. Holothuridea were virtually absent from oysters in both tray types in January and April but biomass increased for both in October (Figure 3.4c). The greater biomass in the October PL-tray communities were not mirrored in the abundance data because this increase was the result of the large size of a single individual of Eupentacta quinquesemita. Except in the October PL-tray community, when  56  Gymnolaemata (particularly Bowerbankia gracilis and Schizoporella unicornis) had the greatest biomass (Figure 3.4b), Polychaeta (particularly Platynereis bicanaliculata, Ophiodromus pugettensis and Eulalia quadrioculata) were the greatest contributors to community biomass, making up between 35% and 70% of the total community biomass in each community.  3.3.1.3 Species richness The species richness data was normally distributed and thus did not require log transformation. Species richness of the fouling communities demonstrated seasonal variation, but no differences between tray types were detected on any sampling date (Table 3.2). Species richness in the January communities was lower than that of the April or October communities (Tukey F=4.75, DF=12, p<0.05); species richness in the April and October communities were comparable (Figure 3.2c). Seasonal differences in species richness were detected for 17 of the 21 major taxonomic groups (Table 3.1). For the majority of the groups, species richness was lowest in January and highest in either October (see Figure 3.5a for example) or April (see Figure 3.5b for example) depending on the group. Hence, similar to the results for total abundance, the seasonal trends in the species richness of the major taxonomic groups are reflected in the seasonal trends of total species richness (Figure 3.2c). a  b  c  40  30  20  10  0  log_Gymnolaemata SR (# spp. / sq.m)  100  log_Polychaeta SR (# spp. / sq.m)  log_Malacostraca SR (# spp. / sq.m)  50  80  60  40  20  0 1 Jan  2 Apr  Date  3 Oct  1 Jan  2 Apr  Date  3 Oct  25  20  15  10  5  0 1 Jan  2 Apr  Date  3 Oct  Figure 3.5. Seasonal mean species richness per fouling community (number species per m2) on oysters suspended in PL and PV-trays for (a) Class Malacostraca, (b) Class Polychaeta and (c) Class Gymnolaemata.  Class Gymnolaemata was the only major group with a difference in species richness between tray types (Table 3.1). The ANOVA results showed a greater number of species on oysters in PL-trays in April and October, than in PV-trays (Figure 3.5c). In April and October,  57  the PL-tray communities had 6 species and 3 species, respectively that were not present in any of the PV-tray communities. A significant interaction between date and tray type was present for Turbellaria and Ophiuroidea showed marginal significance (Table 3.1). In the 6 oyster fouling communities (3 sampling dates x 2 tray types), Class Polychaeta had the greatest number of species present, contributing between 28% (October PV-tray) and 44% (January PV-tray and April PV-tray) of the total number of species in each community. Of the total 182 species identified in the oyster fouling communities, 40 species occurred in all 6 of the fouling communities and belonged to the following groups: Polychaeta (18 species), Malacostraca (5 species), Gymnolaemata (4 species), Bivalvia (3 species), Gastropoda (2 species), Foraminifera (2 species), and Maxillapoda (1 species), Asteroidea (1 species), Ascidiacea (1 species), Demospongiae (1 species), Hydrozoa (1 species) and Turbellaria (1 species).  3.3.1.4  Swartz Dominance Index (SDI)  The SDI of the oyster fouling communities demonstrated seasonal variation (ANOVA F(2, 12)=104.68,  p<0.01). The smallest SDI values (highest dominance) were observed in the October  oyster fouling communities (PL-tray 7.3+0.3 SDI/community; PV-tray 6.7+0.3SDI/community) while the largest values (lowest dominance) were observed in the April fouling communities (PLtray 18.0+0.6 SDI/community; PV-tray 15.7+0.9SDI/community (Figure 3.6). SDI values for the January oyster fouling communities were between the values of April and October (Figure 3.6). The SDI values for the April fouling communities were significantly higher than those of the October communities, and with a single exception, the SDI values for the January fouling communities were different from both April and October communities (Tukey F=4.75, DF=12, p<0.05); only the SDI values of the January PV-tray and April PV-tray communities were not different from each other. The low SDI values of the October oyster fouling communities are largely due to a significant increase in the abundance of several species (e.g. Chrysopetalum occidentale, Aorides intermedius and Calanoid sp.) present in the communities on the other dates but in much lower abundance, as well as the presence and large abundance of the gastropod Odostomia columbiana, a species found only in the October communities. The SDI values demonstrated no significant variation in the oyster fouling communities from the different tray types (ANOVA F(1, 12)=1.96, p=0.19) and there was no interaction between date and tray type (ANOVA F(2, 12)=2.44, p=0.13).  58  Dominance ( SDI / community )  20  Tray_Type PL PV  15  10  5  0 1 January  2 April  3 October  n=3  n=3 Date  n=3  Figure 3.6. Seasonal mean dominance per fouling community on oysters suspended in PL and PV-trays (SDI per community + SE).  3.3.1.5 Community analyses Cluster analyses of the species abundance data and the species biomass data were conducted separately and, in general, results support those of the analyses of variance conducted on total (community) and major group abundance and biomass (see section 3.3.1.1 and 3.3.1.2). The dendrogram generated using species abundance data shows that the October communities’ cluster (both tray types) was significantly different (dissimilarity of 0.92) from the cluster consisting of the January and April communities (both tray types) (Figure 3.7). The null hypothesis that these 2 groupings were homogenous could be rejected at p=0.05 (see linkage A, Figure 3.7). January and April communities were most similar to each other (dissimilarity of 0.64, p>0.05) (see linkage B, Figure 3.7). For each sampling date, the species abundances in the oyster fouling communities from each of the 2 tray types were more similar to each other than to the communities from other sampling dates, with no differences between the tray communities on any date (see Figure 3.7, linkages C, D, and E). The results of the cluster analysis using the species biomass data from the oyster fouling communities were essentially the same as those using the species abundance data. The dendrogram illustrates that, based on species biomass, the October communities’ cluster (both tray types) was significantly different from the cluster consisting of all the January and April communities (dissimilarity of 0.74) (Figure 3.8). The null hypothesis that these 2 groupings were homogenous could be rejected at p=0.05 (see Figure 3.8, linkage A). Similar to the results for  59  species abundance, there were no differences between the species biomass in the oyster fouling communities for each of the 2 tray types, and for each sampling date (see Figure 3.8, linkages C, D, and E).  E  October-PL October-PV  A A D  *  January-PL January-PV  B  April-PL  C  April-PV  Dissimilarity  Figure 3.7. Hierarchical cluster dendrogram showing the relative dissimilarities of fouling communities using an unweighted pair-group mean average sort algorithm based on Bray-Curtis dissimilarity index values of species abundance data. Each letter (A through G) is associated with a cluster linkage. * linkage where the non-parametric SIGTREE bootstrap method (200 simulations) detected the paired groups “clusters” were significantly different from each other.  October-PL  C  October-PV  A A  E  *  January-PL January-PV  B  April-PL  D  April-PV  Dissimilarity  Figure 3.8. Hierarchical cluster dendrogram showing the relative dissimilarities of fouling communities using the Bray-Curtis dissimilarity index based on species biomass data. Each letter (A through G) is associated with a cluster linkage. * linkage where the non-parametric SIGTREE bootstrap method (200 simulations) detected the paired groups were significantly different from each other.  60  3.4  DISCUSSION Cultured oysters provide an abundance of available substrate suitable for invertebrate  larvae to settle and attach. In this study, the shells of C. gigas grown in deep-water suspended tray culture supported diverse fouling communities. These communities were found to undergo seasonal variation in abundance and biomass. Seasonal fluctuations in temperature, salinity and chlorophyll (indicative of phytoplankton) are known to affect the growth and reproduction of marine fouling communities (Osman, 1977). These seasonal changes in key variables influence food availability for many fouling organisms and the cultured oysters, as well as organismal reproduction and larval dispersion (Trujillo and Thurman, 2008). In this study, total abundance and biomass of the oyster fouling communities in both tray types varied seasonally. In January, the low temperatures and chlorophyll levels (see Section 2.3.2, Figure 2.10 for water column details) coincided with the low abundance and biomass of fouling communities, suggesting little or no recruitment, reduced growth and organismal die off in the winter (Arakawa, 1990). The fouling communities’ abundance and biomass in April were similar to January; however, the water column data in April would have predicted a shift in organismal productivity, with increases in chlorophyll and TSS, not reflected in the April fouling communities. It is likely that the abundance and biomass of the fouling communities in this study would have peaked in the summer months (July), with the increased plankton and larvae available in the water column in April and possibly in the subsequent months. The increase in total abundance and biomass of fouling organisms in the October communities is not supported by the October water column data. However, during the 3 months these trays were deployed the water column may have had significant plankton and larvae available to these fouling communities and without these data it is difficult to make specific inferences. Seasonality in biomass of shell fouling communities has been documented in the literature for a variety of cultured bivalves. A fouling study in Indonesia on pearl oysters (Pinctada maxima) suspended in panel nets documented changes in fouling biomass over time, due to the cyclic nature of larval settlement (Taylor et al., 1997). LeBlanc et al. (2003) reported that fouling biomass on mussel socks, deployed in Prince Edward Island from May to November, peaked in September/October. Erbland and Ozbay (2008) studied the biomass of fouling communities on oysters (Crassostrea virginica) growing in an oyster reef and reported a greater biomass of fouling collected in August, to that in September or October (oysters deployed the end of June, oysters cleaned every 2 weeks and fouling collected every 4 weeks).  61  There is evidence in the literature that some fouling organisms may preferentially choose particular substrates over another (Dahlem et al., 1984; Callow and Fletcher, 1994; Tyrrell and Byers, 2007; Erbland and Ozbay, 2008), but it is unclear if or why Demospongiae, Holothuridea, Gymnolaemata and Hydrozoa would preferentially choose to occupy and grow (greater biomass) on oysters in PL-trays more than oysters in PV-trays. Because the 4 groups were present on oysters in both tray types, but biomass was greatest on the oysters in the PL-trays, suggests that oyster in the PL-trays supported a more favourable substrate for growth of these species (particularly Halichondria bowerbanki and Bowerbankia gracilis, which are encrusting organisms), not necessarily that the substrate was more favourable for settlement. The total number of species identified in the oyster fouling communities varied across seasons. Similar seasonal variation in species richness was observed for the fouling communities on C. gigas growing on suspended cultch long-lines on the Mediterranean coast: monitored from January to December, where species richness was highest in the spring and lowest in the winter (Mazouni et al., 2001). In all oyster fouling communities Polychaeta contributed between 28% and 44% (34 to 76 species). Murad and Mohammad (1976) also reported a high incidence of polychaetes as fouling species on Pinctada fucata, in Kuwait contributing more than 55% (42 species) of the total number of species present and Arakawa (1990) reported that at least 108 species of polychaetes have been identified on C. gigas suspended raft culture in Japan. In this study, tray type had no affect on species richness of the oyster fouling communities, regardless of date, but examination of the species composition of the communities revealed 15 species that were present only on oysters in the PL-trays and 5 species that were present only on the oysters in the PV-trays. While there is no literature on the fouling communities on oysters cultured within different tray types, the results of this study suggest that the PV-trays discourage the settlement/growth of more invertebrate species on oysters than do the PL-trays. The total number of species documented within all the oyster fouling communities in this study (182 species) was much greater than that documented in similar studies. For example, 10 species were identified in association with C. virginica on an oyster reef in Delaware (Erbland and Ozbay, 2008) and as many as 76 species were identified in fouling communities on the pearl oyster Pinctada fucata, deployed in suspended trays in Kuwait (Murad and Mohammad, 1976). Arakawa (1990) reported that overtime 1964 species have collectively been identified on C. gigas suspended raft culture in Japan. The variation in the species richness of bivalve fouling communities is likely due to both the range of geographic locations and the variation in date and  62  length of time deployed, as well as the methods used for fouling collection and identification of the fouling samples. The increased dominance observed in the October oyster fouling communities in this study is explained by the increased abundance of a few major species: Chrysopetalum occidentale (Polychaeta), Aorides intermedius (Malacostraca), Calanoid sp. (Maxillapoda) and Odostomia columbiana (Gastropoda). The abundance of O. columbiana on oysters is of particular importance because the genus Odostomia is well documented as an ectoparasite to a number of hosts including oysters such as C. virginica and Pecten maximus (Cole, 1951; Cole and Hancock, 1955; Boss and Merril, 1965). Odostomia on P. maximus have been reported as lodging themselves in the margins of the oyster shell, piercing through the body wall and sucking liquid or tissue from the oyster using their buccal pump (Cole and Hancock, 1955). Oysters exposed to Odostomia parasitism were shown by Cole and Hancock (1955) to have malformation to the edge of shell and in severe cases to have damage to the adductor muscle leaving the oyster gaping. In this study, Odostomia were often found in the crevices of the oyster shell, but no noticeable damage was observed to the shell of oysters. The presence of these organisms on oysters, particularly young, small oysters, should be closely monitored by farmers to ensure no significant damage occurs. Cultured oysters may compete with fouling organisms for food, especially if food sources are limited in particular seasons. As filter-feeders, oysters may be competing (exploitation competition) with other filter-feeders, such as, ascidians, polychaetes, bivalves, barnacles or gastropods that target similar-sized particles (2 to 8µm) at comparable filtration rates (1.8 to 3.2L/hr/g (dry weight)) (Lesser et al., 1992; Claereboudt et al., 1994; Ward et al., 1998; Ropert and Goulletquer, 2000; Le Blanc et al., 2002). Cultured oysters may also be subject to interference competition from fouling organisms growing on their shells. This is particularly likely when colonial or encrusting organisms, such as some ascidians, bryozoans and sponges, growing over the oyster such that it is unable to open its valves adequately and is unable to feed adequately (Lesser et al., 1992). A decrease in somatic tissue growth with increased fouling on the shell of scallops (Pecten ziczac) was reported by Lodeiros and Himmelman (2000). Interestingly, however, it has been suggested that the overgrowth of encrusting species on oysters shells may also promote shell growth through stimulation (Murad and Mohammad, 1976; Arakawa, 1990) and an increase in pearl oyster shell length, associated with increased fouling organisms, has been reported (Murad and Mohammad,  63  1976). Increased shell growth without accompanying increase in meat growth is not beneficial to growers. When growing for the half-shell market, growers are trying to create an oyster with greater meat content, not shell content (Lodeiros and Himmelman, 2000). Ascidians (both solitary and colonial) were present throughout this study. As active filter feeders, filtering up to 444 litres/day, ascidians can potentially compete with oysters for food when they occur in high abundance (Fiala-Medioni, 1978; Young, 1989). Ascidians also have a non-selective method of feeding (Rodriguez and Ibarra-Obando, 2008); they may consume large numbers of invertebrate larvae while actively filtering with significant affects on the fouling community structure (Cowden et al., 1984). The adverse effects of Ciona intestinalis on cultured oysters in the Lunenburg Bay area of Nova Scotia were reduced bivalve growth and increasing mortality through competition for food and reduction of water flow (Carver et al., 2003). The 6 ascidian species identified in this study (Corella inflata, Cnemidocarpa finmarkiensis, Bolentia villosa, Distaplia occidentalis, Botrylloides violaceus and Aplidium californicum) were always present in low abundance, but the greatest abundance was reported in October oyster fouling communities. The solitary ascidian C. inflata was the only species present in all fouling communities, and the non-native colonial ascidian, Botrylloides violaceus was the most abundant species (October only). The abundance of B. violaceus on oysters in October may interfere with oyster growth although the majority of the colonies of this species were very young. Polychaetes were present throughout this study with consistently high abundance, species richness and biomass. Of the 62 species of Polychaeta identified, 9 were filter-feeding or suspension feeding species belonging to 4 polychaete families (Serpulidae, Sabellariidae, Sabellidae and Spiororbidae). All 4 families were present in low abundance and biomass and, therefore, were unlikely to aggressively compete for food resources with the oysters they occupied (Lesser et al., 1992; Ropert and Goulletquer, 2000). Species of the bivalve Family Spionidae were present in all fouling communities in this study with their greatest abundance on the October PL-tray oysters. The spionid polychaetes belonging to the genus Polydora were observed burrowing into oyster shells in this study, resulting in the formation of “mud blisters” or burrows in the shell that fill with mud once the worm has vacated (Murad and Mohammad, 1976; Nel et al., 1996). Although the formation of “mud blisters” has not been demonstrated to impact oyster health or meat quality, the visual quality of the oyster shell is reduced and the half-shell market value of the oyster may be affected. The mud blisters also may interfere with shucking (Bower, 2004). While the abundance and biomass estimates for the spionid species in this study  64  were low, it must be recognized that these organisms may often be unobserved when the inner shell is not inspected. Thirteen bivalve species were found in the oyster fouling communities in this study of which 3 species (Mytilus sp., Hiatella arctica and Pododesmus macrochisma) were present in high abundance in October. Mytilus sp. was particularly abundant on the oysters and represents a significant competitor of the oysters for space and food. As filter feeders, Mytilus spp. have a high filtration rate and can filter a range of particle sizes (Lesser et al., 1992). The ability of Mytilus to strongly attach to the oyster shells, using their byssal threads, may lead to the oysters growing together in clumps. Oysters grown in clumps may have reduced value in the half-shell market and if sold as shucked oysters instead, growers face a reduced profit margin. In this study, Mytilus sp. was often observed using hydroids attached to the oysters as substrate for settlement. Okamura (1986) found that the presence of hydroids in fouling communities directly influences the recruitment of Mytilus species which use them for settlement. Sixteen species of bryozoans (Gymnolaemata) were found in the oyster fouling communities in this study; 13 of these species were encrusting or colonial forms which, if sufficiently abundant, could potentially impact oyster growth. Unlike most of the bryozoan species, which were low in abundance, the colonial, filter-feeding Bowerbankia gracilis had high abundance and biomass in the October communities, particularly the fouling communities on oysters in the PL-trays. The presence of B. gracilis in fouling communities in October was also documented in Poland, on PVC settling plates (deployed May to October) (Dziubińska and Janas, 2007). Abundance and biomass of the 2 species of sponge present in the oyster fouling communities (Halichondria bowerbanki and Leucosolenia nautilia) were highest in the October PL-tray community. In October (particularly in PL-trays), the potential competitors B. gracilis and H. bowerbanki were present on the oysters with significant abundance and biomass to exploit and/or interfere with oysters feeding and/or growth. Barnacles are frequently reported as fouling species on cultured bivalves; in particular Balanus species which are reported repeatedly in fouling literature (e.g. Taylor et al., 1997; Loderios and Himmelman, 2000; LeBlanc et al., 2003; Guenther et al., 2006). In April and October, barnacles (Balanus crenatus and Balanus glandula) were present in the oyster fouling communities but with low abundance and biomass. In addition, the barnacles found were small (likely young adults or recently settled juveniles). It is likely that the sampling times employed in  65  this study and the settling times of the barnacles were such that adult barnacles failed to establish themselves thereby minimizing their potential role in the oyster fouling communities studied. Predation, like competition, is likely to play an important role in determining fouling community structure. In Canada, sea stars are a major concern for shellfish growers, particularly those growers with long-line cultch grow-out operations where product is handled rarely and, therefore, is extremely vulnerable to predators (LeBlanc et al., 2003). In BC, the sea stars Pisaster ochraceus and Evasterias troschelli appear to be common predators on oyster cultch lines (pers. obs.). While P. ochraceus was found frequently in this study’s oyster fouling communities (particularly abundant in October), all specimens were juveniles and, therefore, were unlikely to prey on the much larger oysters. In BC, to date, shellfish farmers have seen few of the potentially detrimental effects of invasive, non-native invertebrate species on cultured bivalves, although the presence of nonnative species on aquaculture sites is increasing and their abundance is evident (pers. obs.). Nonnative, introduced species are of increasing concern to community ecologists and bivalve farm operators worldwide due to their potential impact to local biodiversity and to cultured bivalves (Stachowicz et al., 2002; Carver et al., 2003; Anil, 2006; Altman and Whitlatch, 2007). Suspended shellfish aquaculture operations offer suitable habitat for the establishment of introduced species by increasing substrate availability in protected coastal areas. In this study, 8 non-native species were recorded. While 6 of these species occurred in low abundance, the turbellarian Pseudostylochus ostreophagus and the encrusting bryozoan Bowerbankia gracilis were found in high abundance in the April and October oyster fouling communities, respectively. At the present time, the seasonal abundance of these non-native species has an unknown effect on the growth and health of the cultured oyster or on the fouling community composition. When growing for the half-shell market, growers may frequently handle the oysters and trays (oyster tumbling, removing oysters to decrease growing density, cleaning cages) and this handling may become a crucial component in minimizing the future impact of non-native species on oyster farms in BC.  3.5  CONCLUSIONS  Pacific oysters cultured in suspended oyster trays provide a three-dimensional habitat that supports diverse communities and offers refuge from predators. In this study, oysters cultured in suspended trays supported complex invertebrate communities, which support a greater abundance,  66  biomass and richness of species than those reported in comparable fouling studies to date. This study demonstrated that oysters grown in the plastic and polyvinyl trays commonly used in BC’s oyster industry had comparable fouling communities but that seasonal variation occurred in the abundance, biomass and composition of the oyster fouling communities regardless of tray type. For most marine invertebrates, spring, summer and early fall are the periods of greatest recruitment and growth. Handling of the oysters and trays is likely important in reducing fouling biomass and, given the results of this study, should be performed in late-summer or early-fall to avoid ongoing establishment and growth of large competitive species (e.g. Balanus spp. and Mytilus spp.), to reduce the growth of colonial and encrusting species (e.g. H. bowerbankia and B. gracilis) and to avoid parasitic species (e.g. Odostomia columbiana). Further research should include: the effects of fouling community structure on oyster growth, optimal times for seeding or product deployment, cleaning/handling schedules, as well as comparing oyster fouling communities between farm sites. An interesting addition to an in situ study, such as the one presented in this paper, would be the monitoring of the local larval supply (using plankton sampling and a diversity of settlement panels, including oyster shell) to determine the composition of the larval populations in the water column, the larvae that settle and the preferred substrates. Consistent, long-term monitoring of bivalve fouling communities would provide invaluable data on the distribution and abundance (recruitment) of critical species, including non-native, invasive species.  67  LITERATURE CITED Altman, S., Whitlatch, R.B., 2007. Effects of small-scale disturbance on invasion success in marine communities. Journal of Experimental Marine Biology and Ecology 342: 15-29. Anil, A.C., 2006. A perspective of marine bioinvasion. In multiple dimensions of global environmental change, TERI Press, India. Arakawa, K.Y., 1990. Competitors and fouling organisms in the hanging culture of the Pacific oyster, Crassostrea gigas (Thundberg). Marine and Freshwater Behaviour and Physiology 17: 67-94. Barbour, M.T., Gerritsen, J., 1998. Subsampling of benthic samples: A defense of the fixed-count method. Journal of the North American Benthological Society 15: 386-391. Boss, K.J., Merrill, A.S., 1965. Degree of host specificity in two species of Odostomia (Pyramellidae: Gastropoda). Journal of Molluscan Studies 36: 349-355. Bower, S.M., 2004. Synopsis of Infectious diseases and parasites of commercially exploited shellfish: shell-boring polychaetes of oysters. http://www.pac.dfompo.gc.ca/science/species-especes/shellfish-coquillages/diseases-maladies/pages/sbpoyeng.htm. Bray, J.R., Curtis, J.T., 1957. An ordination of the upland forest communities of southwestern Wisconsin. Ecological Monographs 27: 325-349. Callow, M.E., Fletcher, R.L., 1994. The influence of low surface energy materials on bioadhesion- A review. International Biodeterioration and Biodegradation 34: 333-348. Carver, C.E., Chisholm, A., Mallet, A.L., 2003. Strategies to mitigate the impact of Ciona intestinalis (l.) biofouling on shellfish production. Journal of Shellfish Research 22: 621631. Claereboudt, M.R., Bureau, D., Côté, J.. Himmelman, J.H., 1994. Fouling development and its effect on the growth of juvenile giant scallops (Placopecten magellanicus) in suspended culture. Aquaculture 121: 327-342. Cole, H.A., 1951. An Odostomia attacking oysters. Nature 168: 953-954. Cole, H.A., Hancock, D.A., 1955. Odostomia as a pest of oysters and mussels. Journal of Marine Biology 34: 25-31. Cowden, C., Young, C.M., Chia, F.S., 1984. Differential predation on marine invertebrate larvae by two benthic predators. Marine Ecology Progress Series 14: 145-149. Dahlem, C., Moran, P.J., Grants, T.R., 1984. Larval settlement of marine sessile invertebrates on surfaces of different colour and position. Ocean Science and Engineering 9: 225–236.  68  Dalby, J.E., Young, C.M., 1993. Variable effects of ascidian competitors on oysters in a Florida epifaunal community. Journal of Experimental Marine Biology and Ecology 167: 47-57. DFO, 2005. Juan de Fuca Strait to Strait of Georgia #3462. DFO, Canadian Hydrographic Service. DFO, 2006. Assessing Habitat Risks Associated with Bivalve Aquaculture in the Marine Environment. DFO, Canadian Science Advisory Secretariat, Science Advisory Report 2006/005. Dziubińska, A., Janas, U., 2007. Submerged objects – a nice place to live and develop. Succession of fouling communities in the Gulf of Gdańsk, Southern Baltic. International Journal of Oceanography and Hydrobiology 36: 65-78. Erbland, P.J., Ozbay, G., 2008. A comparison of the macrofaunal communities inhabiting a Crassostrea virginica oyster reef and oyster aquaculture gear in Indian River Bay, Delaware. Journal of Shellfish Research 27: 757-768. Fiala-Medioni, A., 1978. Filter-feeding ethology of benthic invertebrates (Ascidians). IV. Pumping rate, filtration rate, filtration efficiency. Marine Biology 48: 243-249. Gray, J.S., 1997. Marine biodiversity: patterns, threats and conservation needs. Biodiversity and Conservation 6: 153-175. Guenther, J., Southgate, P.C., de Nys, R., 2006. The effect of shell size on accumulation of fouling organisms on the Akoya pearl oyster Pinctada fucata (Gould). Aquaculture 253: 366-373. Hintze, J. 2007. NCSS and GESS. NCSS, LLC. Kaysville, Utah. www.ncss.com. LeBlanc, A.R., Landry, T., and Miron, G. 2003. Identification of fouling organisms covering mussel lines and impacts of a common defouling method on the abundance of foulers in Tracadie Bay, Prince Edward Island. Canadian Technical Report of Fisheries and Aquatic Sciences 2477: 1-26. Lesser, M.P., Shumway, S.E., Cucci, T., Smith, J., 1992. Impact of fouling organisms on mussel rope culture: Interspecific competition for food among suspension-feeding invertebrates. Journal of Experimental Marine Biology and Ecology 165: 91-102. Lodeiros, C., Garcia, N., 2004. The use of sea urchins to control fouling during suspended culture of bivalves. Aquaculture 231: 293-298. Lodeiros, C.J.M., Himmelman, J.H., 2000. Identification of factors affecting growth and survival of the tropical scallop Euvola (Pecten) ziczac in the Golfo de Cariaco, Venezuela. Aquaculture 182: 91-114.  69  Mazouni, N., Gaertner, J.C., Deslous-Paoli, J.M., 2001. Composition of biofouling communities on suspended oyster cultures: An in situ study of their interactions with the water column. Marine Ecology Progress Series 214: 93-102. McKindsey, C.W., Anderson, M.R., Barnes, P., Courtenay, S., Landry, T., Skinner, M., 2006. Effects of shellfish aquaculture on fish habitat. DFO, Canadian Science Advisory Secretariat, Research Document 2006/011. Murad, B.M., Mohammad, F.L.S., 1976. Relationship between biofouling and growth of the pearl oyster Pinctada fucata (Gould) in Kuwiat, Arabian Gulf. Hydrobiologia 51: 129-138. Nel, R., Coetzee, P.S., Van Niekerk, G., 1996. The evaluation of two treatments to reduce mud worm (Polydora hoplura Claparéde) infestation in commercially reared oysters (Crassostrea gigas Thundberg). Aquaculture 141: 31-39. Nemec, A.F.L., Brinkhurst, R.O., 1988. Using the bootstrap to assess statistical significance in the cluster analysis of species abundance data. Canadian Journal of Fisheries and Aquatic Science 45: 965-970. Okamura, B., 1986. Formation and disruption of aggregations of Mytilus edulis in the fouling community of San Francisco Bay, California. Marine Ecology Progress Series 38: 275-282. Osman, R.W., 1977. The establishment and development of a marine epifaunal community. Ecological Monographs 47: 37-63. Rodriguez, L.F., Ibarra-Obando, S.E., 2008. Cover and colonization of commercial oyster (Crassostrea gigas) shells by fouling organisms in San Quintin, Mexico. Journal of Shellfish Research 27: 337-343. Ropert, M., Golletquer, P., 2000. Comparative physiological energetics of two suspension feeders: Polychaete annelid Lanice conchilega (Pallas 1766) and cupped oyster Crassostrea gigas (Thundberg 1795). Aquaculture 181: 171-189. Ross, K.A., Thorpe, J.P., Brand, A.R., 2004. Biological control of fouling in suspended scallop cultivation. Aquaculture 229: 99-116. Sneath, P.H.A., Sokal, R.R., 1973. “Numerical taxonomy: The principles and practice of numerical classification”. W.H. Freeman, San Francisco, 573 pp. Stachowicz, J.J., Fried, H., Osman, R.W., Whitlatch, R.B., 2002. Biodiversity, invasion resistance and marine ecosystem function: Reconciling pattern and process. Ecology 83: 2575-2590. Swartz, R.C., 1978. Techniques for sampling and analyzing the marine macrobenthos. U.S. Environmental Protection Agency (EPA), Doc. EPA-600/3-78-030, EPA, Corvallis Oregon. 27p.  70  Taylor, J.J., Southgate, P.C., Rose, R.A., 1997. Fouling animals and their effect on the growth of silver-lip pearl oysters, Pinctada maxima (Jameson) in suspended culture. Aquaculture 153: 31-40. Trujillo, A.P., Thurman, H.V., 2008. Essentials of oceanography. 9th ed. Pearson Prentice Hall, New Jersey. Tyrrell, M.C., Byers, J.E., 2007. Do artificial substrates favour nonindigenous fouling species over native species? Journal of Experimental Marine Biology and Ecology 342: 54-60. Wahl, M., 1989. Marine epibiosis. I. Fouling and biofouling: Some basic aspects. Marine Ecology Progress Series 58: 175-189. Walters, L.J., Wethey, D.S., 1991. Settlement, refugees and adult body form in colonial marine invertebrates: A field experiment. Biological Bulletin 180: 112-118. Ward, J.E., Levinton, J.S., Shumway, S.E., Cucci, T., 1998. Particle sorting in bivalves: In vivo determination of the pallial organs of selection. Marine Biology 131: 283-292. Woodin, S.A., 1991. Recruitment of infauna: Positive or negative cues. American Zoologist 31: 797-807. Young, C.M., 1989. Larval depletion by ascidians has little effect on settlement of epifauna. Marine Biology 102: 481-489.  71  CHAPTER 4: Summary and conclusions Suspended oyster aquaculture operations support diverse invertebrate fouling communities by providing an abundance of biotic (oysters) and abiotic or artificial (oyster trays, rafts, ropes and buoys) substrates for the settlement of sessile invertebrates and the associated mobile predators. The importance of these fouling communities to oyster growers is two-fold. Firstly, they increase the labour involved in growing a marketable product and, secondly they decrease the market price attainable. This study examines the invertebrate fouling communities on suspended Pacific oyster trays (Chapter 2) and on the oysters themselves (Chapter 3). The composition of the fouling communities was examined for seasonal variation, with samples collected every 3 months (excluding July oyster fouling) over the course of 1 year. In addition, fouling communities on trays and oysters were compared between the plastic trays (PL) and the polyvinyl trays (PV). In general the 2 substrate types examined, trays and oysters, supported fouling communities of similar species composition. Except for the major taxonomic group Actinopterygii, the same groups were present in both the tray and oyster communities. Considering all identified species in the fouling communities on trays (194 species) and oysters (182 species) the 2 community types had 163 species in common. The tray communities had 31 species that were not present in the oyster communities and the oyster communities had 19 species that were not present in the tray communities.  4.1 Seasonal variation in fouling community composition Both the tray and oyster fouling communities had seasonal changes evident in abundance, biomass and species richness that correlated with seasonal changes in the local water conditions, particularly the abundance of plankton measured as chlorophyll. Similar relationships between fouling community composition and water column variables have been documented in other fouling studies (Osman, 1977; Oshurkov, 1992; Okamura, 1986; Nair, 1967; Turner and Todd, 1993). Unlike the tray fouling communities, the oyster fouling communities were not examined in July and, as a result, only limited seasonal comparisons can be made between the tray and oyster fouling communities. Tray fouling communities (both tray types) had the greatest abundance, biomass and species richness (PL-tray only) in July. In contrast, oyster fouling communities had the greatest abundance and biomass in October and species richness was greatest in the April and October communities. The greatest contributors to abundance, biomass and species richness in the trays were Polychaeta, Malacostraca, Bivalvia and Gymnolaemata.  72  However, in the oyster fouling communities only Polychaeta contributed greatly to abundance, biomass and species richness, although the Gymnolaemata did contribute to biomass of the oyster fouling in the October PL-trays. Differences in tray (abiotic) and oyster (biotic) fouling communities may indicate the documented trend that some organisms exhibit substrate preference (Dahlem et al., 1984; Walters, 1992; Callow and Fletcher, 1994), with the preferred substrates providing, for example, protection from visual predators (tray) or protection from small motile invertebrates (oyster crevices) (McKindsey et al., 2006). If oyster fouling was examined in July, as it was for tray fouling, major groups such as Malacostraca and Bivalvia may have been greater contributors.  4.2 Tray-type and fouling community composition The oyster aquaculture industry has developed measures to increase the potential profit margin for growers, including the introduction of a new PV-tray that stacks 10 trays high compared to the 7 trays high in the conventional PL-tray, thereby increasing the stocking density of the oyster stack by 43%. It also was anticipated that the new PV-tray would reduce fouling in comparison with the PL-tray because the latter tray type offered larger solid surfaces within each tray for settlement. In addition, it was thought that larvae may choose not to settle on the PV-trays (Dahlem et al., 1984; Woodin, 1991; Walters and Wethey, 1991; Callow and Fletcher, 1994). This study demonstrated that PV-trays failed to reduce the overall abundance or overall biomass of the fouling communities on the trays or on the oysters. However, species richness was reduced on the PV-trays. When the abundance, biomass and species richness of the major taxonomic groups present in the fouling communities were examined, some differences between the 2 tray types were detected for both the tray and oyster fouling communities. For example, species richness in the tray fouling communities was greater on PL-trays than PV-trays for Polychaeta, Asteroidea and Anthozoa and, species richness in the oyster fouling communities was greater in PL-trays than PV-trays for Holothuridea, Demospongiae, Gymnolaemata and Hydrozoa.  4.3 Research implications This study did not suggest that PV-trays significantly reduce fouling on either the trays or the oysters. Some major groups did have less abundance, biomass and/or species richness on PVtrays and their oysters. The reduction of particular groups on the PV-trays and on the oysters may  73  be of particular importance if they include species that are potential competitors to the oysters. For example, a reduction in the abundance or biomass of groups such as, Polychaeta, Bivalvia, Gastropoda and Anthozoa on PV-trays may positively impact the growth of oysters. Because there is no doubt that the use of the PV-trays increases the number of oysters that can be grown on an oyster raft, and they do reduce some key fouling organisms, I would state that the PV-tray is generally more suitable to oyster growers, however, I believe there is considerable room for improvement and because the trays are costly farmers, they should weigh the cost-benefits of such a large investment. The greatest draw back to the PV-tray, is the use of the plastic liner in the bottom of the trays, which can trap more fouling between the tray and liner than would otherwise accumulate. Fouling communities vary seasonally, with the greatest amount of fouling (biomass) on both trays and oysters occurring through the summer and early fall. To avoid fouling, growers should consider adjusting farming practices, such as seeding times and antifouling techniques, accordingly. Farmers should consider putting new product or seed at a grow-out site between mid-fall and late-winter, after most invertebrate larval settlement has taken place to allow oysters significant time to grow before they are exposed to extensive fouling recruitment and subsequently antifouling treatment or tumbling. To maximize cost-effectiveness growers should perform any antifouling techniques including oyster tumbling, in early-summer after initial spring settlement and before substantial growth and early-fall after further settlement and growth.  4.4 Future research The introduction of aquaculture operations, and associated substrates, into BC’s coastal waters undoubtedly provides additional substrate for fouling organisms. The impact of these fouling organisms on bivalve culture may be mitigated through the development of an oyster tray that reduces fouling. Development of a new tray would require additional investigation into substrates, or substances, that discourage potentially harmful species, such as mussels and ascidians from settling. These investigations would need to include research aimed at an improved understanding of larval recruitment, settlement cues and larval substrate selectivity for harmful fouling species. Settling, or fouling, plates are used often to examine species-specific settling behaviour and fouling community development and would be useful for studying fouling species of interest to BC’s bivalve farmers. Both abiotic and biotic substrates can be treated with mechanical, chemical and biological methods to control fouling communities and, although a  74  number of examples of these methods have been tested, the majority are limited to specific species and localities and many have been tested only in the laboratory. Additional research into these methods, that is directly applicable to fouling organisms of relevance to BC’s oyster aquaculture industry, is required. Fouling communities associated with suspended oyster aquaculture provide an excellent opportunity to study hard-substrate communities, species-specific recruitment and settlement patterns, biological interactions (competition and predation) and marine non-native, particularly invasive species (abundance, distribution, morphology and life histories). Future research should utilize these facilities to further our understanding of fouling species of economic and ecological importance. For example, invasive species may impact both economic and ecological processes through competition with cultured bivalves and through alteration of local coastal marine biodiversity and ecosystem function.  75  LITERATURE CITED Callow, M.E., Fletcher, R.L., 1994. The influence of low surface energy materials on bioadhesion- A review. International Biodeterioration and Biodegradation 34: 333-348. Dahlem, C., Moran, P.J., Grants, T.R., 1984. Larval settlement of marine sessile invertebrates on surfaces of different colour and position. Ocean Science and Engineering 9: 225–236. McKindsey, C.W., Anderson, M.R., Barnes, P., Courtenay, S., Landry, T., Skinner, M., 2006. Effects of shellfish aquaculture on fish habitat. DFO, Canadian Science Advisory Secretariat, Research Document 2006/011. Nair, N.U., 1967. The settlement and growth of major fouling organisms in Cochin Harbour. Hydrobiologia 30: 503-512. Okamura, B., 1986. Formation and disruption of aggregations of Mytilus edulis in the fouling community of San Francisco Bay, California. Marine Ecology Progress Series 38: 275-282. Oshurkov, V.V., 1992. Succession and climax in some fouling communities. Biofouling 6: 1-12. Osman, R.W., 1977. The establishment and development of a marine epifaunal community. Ecological Monographs 47: 37-63. Turner S.J., Todd, C.D., 1993. The early development of epifaunal assemblages on artificial substrata at two intertidal sites on an exposed rocky shoe in St. Andrews Bay, N.E. Scotland. Journal of Experimental Marine Biology and Ecology 166: 251-272. Walters, L.J., 1992. Post-settlement success of the absorbent bryozoan Bugula neritina (L.): the importance of structural complexity. Journal of Experimental Marine Biology and Ecology 164: 55-71. Walters, L.J., Wethey, D.S., 1991. Settlement, refugees and adult body form in colonial marine invertebrates: A field experiment. Biological Bulletin 180: 112-118. Woodin, S.A., 1991. Recruitment of infauna: Positive or negative cues. American Zoologist 31: 797-807.  76  

Cite

Citation Scheme:

        

Citations by CSL (citeproc-js)

Usage Statistics

Share

Embed

Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                        
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            src="{[{embed.src}]}"
                            data-item="{[{embed.item}]}"
                            data-collection="{[{embed.collection}]}"
                            data-metadata="{[{embed.showMetadata}]}"
                            data-width="{[{embed.width}]}"
                            async >
                            </script>
                            </div>
                        
                    
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:
http://iiif.library.ubc.ca/presentation/dsp.24.1-0069194/manifest

Comment

Related Items