Re‐Establishment of the Native Oyster, Ostrea Conchaphila, in Netarts Bay, Oregon, USA

AN ABSTRACT OF THE THESIS OF

Pamela Emily Archer for the degree of Master of Science in Marine Resource Management presented on August 6, 2008.

Title: Re‐establishment of the Native Oyster, Ostrea conchaphila, in Netarts Bay, Oregon, USA.

Abstract approved:

Jessica A. Miller

Olympia oysters, Ostrea conchaphila, were once common along the west coast of North America. A popular delicacy, native oyster populations began to decline in the late 1800’s due to over‐harvest, degraded water quality, and habitat loss. Interest in re‐establishing the native oyster in a small Oregon estuary, Netarts Bay, culminated in a partnership among The Nature Conservancy, the National Oceanic and Atmospheric Administration, the Oregon Watershed Enhancement Board, and Oregon State University. This study was designed to assess the re‐ establishment progress of the Olympia oyster restoration in Netarts Bay along with subsequent impacts of the restoration on eelgrass (Zostera marina), an important estuarine species.

Two brood years (2005 & 2006) of cultch, consisting of O. conchaphila set on clean Crassostrea gigas shell substrate, were outplanted within an extensive, relatively uniform eelgrass bed. Cultch was placed in two experimental locations to determine the effect of cultch cover on native oyster survival, growth, and eelgrass abundance. The percent cover of cultch varied among treatments: “control” (no cultch), “low” (4% cultch cover), “medium” (11% cultch), and “high” (19% cultch). Research objectives were: (1) determination of O. conchaphila density, growth, and reproduction; and (2) quantification of the response of Z. marina abundance and reproduction to cultch cover. Results from 2007 demonstrated that Olympia oysters were capable of growth, reproduction, and recruitment within their former habitat. Cultch cover within treatments did not change throughout the summer and there was minimal shell export out of the experimental location. Oyster size increased from March‐September, 2007: the mean size of the 2005 brood year increased by 10.5 mm, while the 2006 brood year increased by 16.2 mm. Sperm and larvae were found in individuals from both brood years, indicating that oysters were reproductively active. Declines in eelgrass mean percent leaf cover and shoot density were observed with increasing cultch cover. The mean eelgrass percent leaf cover was 15‐22% lower and shoot density was 27‐36% lower in high treatment (19% cultch) plots than in control plots. There were no discernable patterns in the eelgrass response variables of flowering shoot count, blade length, or blade width. The medium treatment (11% cultch), in which oyster densities were statistically similar to the high treatment (19% cultch), did not have statistically significant impacts on eelgrass percent cover or shoot density. We recommend continued testing of the medium treatment (11% cultch), as well as other cultch densities, such as a 50% cultch treatment. Additional monitoring will be needed to determine what, if any, long‐term impacts occur to the eelgrass bed. We also recommend long‐term monitoring of both oysters and eelgrass beds to detect any additional changes at the re‐establishment site.

Re‐establishment of the Native Oyster, Ostrea conchaphila, in Netarts Bay, Oregon, USA.

by Pamela Emily Archer

A THESIS

Submitted to

Oregon State University

in partial fulfillment of the requirements for the degree of

Master of Science

Presented August 6, 2008 Commencement June 2009 Master of Science thesis of Pamela Emily Archer presented on August 6, 2008.

APPROVED:

Major Professor, representing Marine Resource Management

Dean of the College of Oceanic and Atmospheric Sciences

Dean of the Graduate School

I understand that my thesis will become part of the permanent collection of Oregon State University libraries. My signature below authorizes release of my thesis to any reader upon request.

Pamela Emily Archer, Author

ACKNOWLEDGEMENTS

First, I would like to thank Dr. Jessica Miller and Dick Vander Schaaf for allowing me to partake in the initial restoration of Oregon’s native oyster. Their support and guidance inspired me to expand my knowledge and test my boundaries. I would like to thank Dr. Anthony D’Andrea and the College of Oceanic and Atmospheric Sciences for their in‐kind support of my research. Dr. Ralph Garono provided me with my first graduate research experience, and I am indebted to him for that.

I would like to thank Dr. Michael Harte and the Marine Resource Management Program at Oregon State University for their assistance and encouragement. I am very appreciative of Dr. D’Andrea and Dr. Robert Wheatcroft for including me into their lab group. Dr. Hal Batcheldor and Dr. Mark Needham each provided welcome recommendations and advice.

The National Oceanic and Atmospheric Administration’s West Coast Native Oyster Working Group is a unique organization of inspired intellectuals who provided me with feedback and encouragement. I was also assisted by the faculty and staff at Coastal Oregon Marine Experimental Station and the Hatfield Marine Science Center.

I would like to thank the multiple volunteers who helped complete field work during this project: Robbie & Daniel Wisdom, Stefanie Gera, Abby Nickels, Rebecca Tully, Brent Matteson, Marisa Litz, Summer Peterman, A. Miller Henderson, and many others. The oyster group at Netarts Bay also provided assistance and guidance: Mark Wittwer, John Johnson, Sue Cudd and the Whiskey Creek Shellfish Hatchery, Alan Barton, David Stick, and Dr. Chris Langdon.

I would like to thank Rhea Sanders and Brent Matteson for providing reviews of this work, and my sister, Stephanie Ann Archer, who assisted with my library research. Many other friends, family members, and colleagues provided me with encouragement and advice, and I would not have been able to complete this work without their valued, unconditional support.

CONTRIBUTION OF AUTHORS

Dr. Jessica Miller

Dr. Miller assisted in the development of the re‐establishment project and the experimental design. Dr. Miller provided academic and research support, helped with field work and overall project logistics. As my major professor, Dr. Miller provided insight and feedback about the project and the development of my research questions. She provided helpful guidance with data analysis, writing, and revising.

Dr. Anthony D’Andrea

Dr. D’Andrea assisted with data analysis, writing, and revising, and helped provide an ecological framework to the writing.

Mr. Dick Vander Schaaf

Mr. Vander Schaaf developed the Olympia oyster re‐establishment plan for Netarts Bay, Oregon. He coordinated the grants, funding, outreach, project logistics, set up the re‐ establishment site, and coordinated the oyster outplantings. Mr. Vander Schaaf also assisted with revisions.

Mr. David Stick

Mr. Stick assisted with oyster DNA extraction and performed the PCR analyses. He also co‐authored the sections on DNA analyses and provided feedback on oyster growth patterns. TABLE OF CONTENTS

Page

Chapter 1: General Introduction ...... 1

The decline of Ostrea conchaphila and current re‐establishment efforts ...... 1

Project background ...... 3

Ecology of Ostrea conchaphila ...... 4

Oyster nutrition ...... 4

Life history & reproduction of O. conchaphila...... 5

Reef structures ...... 5

Ecology of Zostera marina ...... 7

Distribution ...... 7

Eelgrass and essential fish habitat ...... 8

Restoration methods ...... 9

Chapter 2: Ostrea conchaphila re‐establishment ...... 10

Introduction ...... 10

Methods ...... 13

Site description ...... 13

Oyster size & growth...... 16

Oyster reproduction ...... 17

Oyster density ...... 18

Site characterization ...... 18

Eelgrass response to cultch treatments ...... 19

Statistical analyses ...... 19

Results ...... 20

Oyster size and growth ...... 20 TABLE OF CONTENTS, Cont'd

Oyster reproduction ...... 23

Oyster density ...... 26

Site characterization ...... 27

Eelgrass response to cultch treatments ...... 31

Discussion ...... 34

Re‐establishment of oysters within Netarts Bay ...... 34

Oyster density and reef structures ...... 35

Interactions between cultch and eelgrass ...... 36

Additional ecological interactions ...... 37

Recommendations ...... 38

Future research ...... 39

Conclusions ...... 40

Chapter 3: Restoration in Practice ...... 42

Defining long term restoration goals & a reference system ...... 42

Identifying limiting factors ...... 43

Water quality ...... 44

Predators, parasites, and pathogens ...... 44

Aquaculture ...... 45

Incorporating science into the design and monitoring of the project ...... 45

Policy: Long term management of Olympia oysters, eelgrass, and EFH ...... 47

Public involvement and restoration inertia ...... 48

Conclusions ...... 50

Chapter 4: Concluding thoughts ...... 51

Bibliography ...... 53

Appendices ...... 60 LIST OF FIGURES

Figure Page

Figure 1. Netarts Bay, Oregon, USA...... 14 Figure 2. Shell cultch comprised of O. conchaphila spat on C. gigas shells...... 15 Figure 3. Map of experimental locations...... 15 Figure 4. Photo comparison of O. conchaphila to C. gigas...... 16 Figure 5. Oyster monthly mean size in 2007...... 21 Figure 6. Mean monthly oyster growth by treatment...... 21 Figure 7. Size distribution of 2005 brood year...... 22 Figure 8. Size distribution of 2006 brood year...... 22 Figure 9. Photo of September recruitment event...... 24 Figure 10. PCR assay for O. conchaphila...... 25 Figure 11. PCR assay for C. gigas...... 25 Figure 12. Olympia oyster density as compared to cultch treatment...... 26 Figure 13. Mean eelgrass percent cover...... 28 Figure 14. Mean eelgrass shoot density...... 28 Figure 15. Mean proportion eelgrass flowering shoots of total shoots...... 28 Figure 16. U. pugettensis burrow hole density...... 29 Figure 17. Mean macroalgae percent cover by treatment...... 31

LIST OF TABLES

Table Page

Table 1. Netarts Bay sampling regime, Summer 2007...... 16 Table 2. Oyster reproductive tissue...... 24 Table 3. Eelgrass dynamics in control plots for locations A and B...... 27 Table 4. ANOVA on rank‐transformed values of macroalgae percent cover...... 31 Table 5. ANOVA of ArcSin transformed values of eelgrass percent cover...... 33 Table 6. ANOVA of log transformed values of eelgrass shoot count m‐2...... 33 Table 7. ANOVA of log‐transformed values of percent flowering shoots...... 33

LIST OF APPENDICES

Appendix Page

Appendix A. ANOVA tables for 2005 oyster shell length...... 60 Appendix B. Table of oyster reproductive activity by month...... 60 Appendix C. ANOVA table for oyster density...... 60 Appendix D. Mean daily temperatures at experimental site...... 61 Appendix E. Salinity measured at site...... 61 Appendix F. ANOVA details for eelgrass parameters...... 62 Appendix G. Mean macroalage percent cover...... 62 Appendix H. Table of means for eelgrass parameters of percent cover and shoots m‐2...... 62 Appendix I. Graphs of eelgrass blade length...... 63 Appendix J. ANOVA table for eelgrass blade length...... 63 Appendix K. Graphs of eelgrass blade width...... 63 Appendix L. ANOVA table for eelgrass blade width...... 64

DEDICATION

For my mother, Kathryn Ann Leyes,

and

my father, Thomas James Archer. 1

Chapter 1: General Introduction This thesis represents research conducted during the first year of an effort to re‐ establish native oysters along the Oregon Coast. The first chapter provides information on the re‐establishment project, the biology and ecology Olympia oysters, and potential ecosystem effects of the re‐establishment, with a specific focus on eelgrass. The second chapter presents the body of scientific work on the project to be submitted to the journal Restoration Ecology. The third chapter synthesizes the restoration efforts for restoration practitioners and expands on recommendations for future management and restoration of Olympia oysters. The fourth and final chapter provides a conclusion to the thesis as a whole.

The decline of Ostrea conchaphila and current reestablishment efforts The immense popularity of the Olympia oyster, Ostrea conchaphila (= Ostrea lurida = Ostreola conchaphila, Carpenter 1857), led to its demise but may also motivate the re‐ establishment of this native shellfish. Native oysters had been harvested for millennia on the North American West Coast, evidenced by large shell middens with 3000‐4000 year‐old Olympia oyster shells found on estuary shorelines (Cook et al. 1998, Gordon et al. 2001). Two hundred years ago, Olympia oysters were relatively abundant (Baker 1995). When eastern settlers first arrived on the California coast, they described Olympia oysters as a plentiful and delectable with a slight coppery taste (Gordon et al. 2001). San Francisco Bay’s stocks were in high demand and were rapidly depleted shortly after the discovery of gold at Sutter’s Mill in 1849, (Gordon et al. 2001). Consequently, San Francisco began to import large quantities of Olympia oysters from other west coast estuaries throughout the 1850s and 1860s (Kirby 2004). Shiploads of oysters were removed from estuaries such as Humboldt Bay, California, Coos and Yaquina Bays within Oregon, and from Willapa Bay to Puget Sound, Washington. Oyster harvest formed a critical base of regional economics for many towns such as Oysterville, Washington (Steele 1957, Espy 1977).

Olympia oysters are known for their distinct coppery‐metallic taste and are coveted by oyster connoisseurs, who fondly refer to them as “Olys.” The increase in popularity of the Olympia oyster was followed by a rapid population decline within west coast estuaries, with peak harvests of the Olympia oyster ended in the late 1890s (Baker 1995, Kirby 2004). Later attempts to revive some fisheries were met with little success (Baker 1995 and references 2 therein). Harvest techniques, such as dredging, were effective at removing large amounts of oysters and shell but disturbed benthic communities and suspended sediment, which lowered water quality for the oyster. Dredging, along with deep water tonging, helped oystermen collect even the most elusive oysters. The removal of millions of oysters also removed millions of pounds of shell (Townsend 1893), substrate critical for larval settlement.

Oystermen in Puget Sound, Washington, responded to the declines of Olympia oysters and eventually began to cultivate O. conchaphila in diked ponds containing shell substrate (Couch and Hassler 1989). Passage of the Callow Act of 1890 and the Bush Act in 1895 allowed oystermen to purchase the tidelands they farmed as long as the lands remained in oyster production. However, the rest of the O. conchaphila fishery was not regulated until the 1930’s (Baker 1995, Gordon et al. 2001). By this time, populations of O. conchaphila had been severely reduced and had not recovered to previous abundances . This decline was in part due to: lack of substrate for settlement; increases in fine, silty sediments from upper watershed activities; increases in water pollution; and introduced pathogens from Crassostrea virginica and C. gigas (Couch and Hassler 1989, Harris 2004, Ruesink et al. 2005). Oyster growers also noticed a direct effect of sulfite waste liquor, a pollutant released from pulp mills into estuaries, which reduced reproductive success and body weight (Baker 1995). These factors, along with habitat loss through shoreline development and the loss of shell substrate, contributed to the decline of the native oyster population.

Other factors may cause changes in the distribution and abundance of O. conchaphila over time. For instance, O. conchaphila was extinct in Oregon’s Coos Bay before European settlement took place (Baker et al. 1999). In the late 1980’s, O. conchaphila were re‐discovered within deep channels within Coos Bay (Baker et al. 1999). Reasons for resettlement are unknown, although dredging activity may have created water circulation and salinity regimes favorable for the oyster (Baker et al. 1999).

Today, remnant populations of O. conchaphila remain along the west coast but they have not recovered to pre‐1850 abundances (Couch and Hassler 1989, Baker 1995, Ruesink et al. 2005). Harvest of oysters from marine reserves is currently prohibited in California, allowed in Washington only if oysters are greater than 6.35 cm, and completely prohibited in Oregon (California Department of Fish and Game 2008, Oregon Department of Fish and Wildlife 2008,

Washington Department of Fish and Wildlife 2008). There is still consumer demand for Olympia oysters and several boutique oyster growers in Washington continue to culture and market them. In Washington, private aquaculture of O. conchaphila continues today and an oyster stock revitalization plan has been established to rebuild the population (Cook et al. 1998). Other restoration projects have also been developed to re‐establish Olympia oysters along the west coast (see page 9).

Project background The Nature Conservancy of Oregon (TNC) partnered with the National Oceanic and Atmospheric Administration (NOAA), the Oregon Watershed Enhancement Board (OWEB), and the Oregon State University’s Coastal Oregon Marine Experiment Station (COMES) and Molluscan Broodstock Program (MBP) to explore re‐establishment and restoration options for the native Olympia oyster in Netarts Bay, Oregon. TNC’s overall project goal is to re‐establish a self‐sustaining population of native oysters within Netarts Bay. O. conchaphila were reportedly present in the bay around 1900, but by 1918, the “Supply was insufficient for local demands” (Edmondson 1923). A population existed in the southwest corner of the estuary (Marriage 1958); numbers were described as “Extremely low” (Dimick et al. 1941), but no quantitative reports have been found. No documentation was found on the possible extirpation of Olympia oysters from Netarts Bay (Kreag 1979). In the 1990’s, the Oregon Department of Fish and Wildlife (ODFW) attempted to re‐establish Olympia oysters in the mid to upper estuary with an oyster seed source from Willapa Bay, Washington (John Johnson, pers. comm.). A small population of the transplanted oysters remains in Netarts Bay, but no other naturally occurring populations have been found despite much exploration (Dick Vander Schaaf, pers. comm.).

Netarts Bay was designated a conservation estuary under the Oregon Estuary Plan in 1987 to “…Be managed for long‐term uses of renewable resources that do not require major alterations" (Department of Land Conservation and Development 1984). Estuaries with conservation status have limited commercial development, contain valuable natural resources, and restoration projects are encouraged. Netarts Bay is a bar‐built estuary on Oregon’s north coast, south of Tillamook Bay (45°26′0″N 123°56′24″W). The sixth largest estuary in Oregon, Netarts Bay covers approximately 1093 hectares (ha) and drains a watershed area of 3626 ha (Percy et al. 1974). Twelve small creeks provide the only freshwater to the estuary, as there are

4 no major riverine sources (Percy et al. 1974). The estuary is marine‐dominated, has an estimated flushing time of 1.7 tidal cycles, and intertidal areas cover 65% of the estuary (Kreag 1979). Netarts Bay contains the largest eelgrass beds within Oregon, covering 161 ha of tidelands and 175 ha of submerged lands (Stout 1976). Eelgrass beds in Netarts Bay may form Z. marina lakes, described as depressions in the eelgrass bed where water pools at low tide (Stout 1976), reaching depths of ≤10 cm of water (pers. obs.). The bay has commercial oyster beds of C. gigas and is a popular destination for recreational clammers (Stout 1976). Netarts Bay is also home to the invasive green crab Carcinus maenas (Yamada et al. 2005) along with native and non‐native oyster drills. Because of its conservation status, marine‐dominated waters, and broad intertidal areas, Netarts Bay appears to provide quality native and non‐native oyster habitat.

Ecology of Ostrea conchaphila The Olympia oyster is the only oyster native to the Pacific Northwest. Olympia oysters are relatively small, about 3‐5 cm in length. Shells can be completely flat or fluted on the outside, with a characteristic purplish stain from the adductor muscle on the inside (Korringa 1976). Oyster populations have been found within British Columbia, Washington (Willapa Bay, possibly Grays Harbor, Bellingham Bay, Samish Bay, Discovery Bay, Hood Canal, and Puget Sound), Oregon (Netarts Bay, Yaquina Bay, and Coos Bay), California (Humboldt Bay, Tomales Bay, San Francisco Bay, Elkhorn Slough/Monterey Bay, and south beyond Point Conception), and Mexico (Baja California Norte, Baja California Sur) (Fitch 1953, Baker 1995, Groth and Rumrill 2008). These sites, along with other published information on Olympia oysters, can provide useful insight on potential oyster restoration sites and objectives.

Oyster nutrition O. conchaphila are filter feeders which utilize phytoplankton as their primary food source (Baker 1995). Dissolved organic matter and detritus also supplement their diet, as sediment particles provide essential amino acids (Langdon and Newell 1996). Modern, detailed knowledge of Olympia oyster nutrition is unavailable. O. conchaphila has a lower filtration rate than C. gigas (Galtsoff 1932), although the size class of phytoplankton on which O. conchaphila feeds is unknown. An early study on the feeding behavior of both O. conchaphila and C. gigas (then classified as Ostrea gigas) concluded that O. conchaphila were able to filter smaller

5 particles than C. gigas (Elsey 1935). Elsey postulated that because the eggs of O. conchaphila are one‐third larger than those of C. gigas, the ostia through which the eggs pass must also be one‐ third larger. Elsey concluded larger ostia of O. conchaphila may inefficiently filter nanoplankton, a perceived disadvantage compared with C. gigas (Elsey 1935), which could out‐compete the native oyster for food resources. Within the current literature, Elsey (1935) has been interpreted to mean that O. conchaphila filter a larger size class of phytoplankton than C. gigas and therefore fill a different ecological role (Barrett 1963, Couch and Hassler 1989, Baker 1995, Peter‐Contesse and Peabody 2005). Given that Elsey (1935) presents only a qualitative observation that more undigested particles passed through the ostia of O. conchaphila, the conclusion that O. conchaphila and C. gigas consume plankton of different size classes and do not directly compete for food resources is premature. A qualitative laboratory feeding study is needed to quantify the function and filtration capacity of O. conchaphila.

Life history & reproduction of O. conchaphila O. conchaphila are protandrous hermaphrodites which continue to switch reproductive forms throughout their lives (Coe 1932). Spermatogenesis begins 12 weeks after attachment, and oogenesis begins when water temperatures reach 13oC ‐ 16oC for 3‐6 months (Baker 1995). Multiple spawning events within one year are possible. Male oysters expel sperm clusters, termed sperm balls, into the water, which enter the gills of the female oysters. All members of the Ostreniae family, including O. conchaphila, are larviporous, i.e. the eggs are fertilized then retained in the mantle (Harry 1985). In the mantle, the larvae undergo four major stages of development in 10‐14 days, from a zygote to trochophore, D‐hinged larvae, and finally a shelled larva, referred to as a veliger. Up to 250,000 free‐swimming larvae per female are released from the mantle and remain planktonic for two to eight weeks (Baker 1995). Little is known about the pelagic stage of O. conchaphila but individuals settle on hard substrate and grow 35‐45 mm in three years, reaching a mean size of 50 mm, but in rare instances reach 100 mm (Baker 1995, Peter‐Contesse and Peabody 2005). Little growth occurs after the fourth year (Baker 1995). The maximum reported size is 75 mm (Hertlein 1959), but a maximum age has not been reported (Couch and Hassler 1989).

Reef structures Olympia oysters are primarily an intertidal species, found from ‐10 to +2 m relative to mean low water (MLW) (Baker et al. 1999). O. conchaphila prefers waters with an average

6 salinity of 25 (Baker 1995). Oyster larvae require hard substrate to anchor themselves, although substrate type and size varies. Olympia oysters have been found on rocky rip‐rap, dock pilings, gravelly areas, cultured C. gigas, shell rubble, intertidal trees, and in deep channels (Groth and Rumrill 2008).

Historical accounts and descriptions of Olympia oyster reef structures are rare in peer‐ reviewed literature, although qualitative descriptions provide some insight as to how reefs may have existed. Most information is available from government fish and game reports (Fitch 1953, Marriage 1958, Couch and Hassler 1989) and cultural accounts (Steele 1957, Espy 1977, Gordon et al. 2001). Advantages of reef structures provide for the Eastern oyster indicate reef structures are likely advantageous for the survival and propagation of the Olympia oyster (Lenihan 1999, Breitburg et al. 2000, Coen and Luckenbach 2000). It is widely recognized that Olympia oysters form reef aggregates, comprised of patchy clumps of oysters and substrate (Baker 1995). The reef aggregates form at mid‐tidal elevation, between higher elevation eelgrass (Zostera marina) beds and lower elevation non‐vegetated mudflats, but have been found anywhere from 0 to ‐71 m (Hertlein 1959). Dimick (1941) described the native oyster “In great supply on the shoals throughout [Yaquina] Bay,” and Fitch (1953) notes, “In the Pacific Northwest, it is found in beds on the surface of mudflats and gravel bars near the mouths of rivers or streams…attached to the shells of previous generations of oysters or any firm surface that will hold it out of the mud.” Today, oysters found in central California occur naturally as “Clumps…defined as congregations of hard substrate‐creating organisms covering >0.5 m2 of the bottom or as isolated individuals attached to hard surfaces in the intertidal” (Heiman et al. 2008).

It is also possible O. conchaphila reef aggregates included areas of eelgrass. The discovery of Pleistocene fossils in northern California indicated O. conchaphila occurred in “Isolated clumps surrounded by slightly shelly mud matrix” (Miller and Morrison 1988). Eelgrass (Zostera spp.), was also found in this fossilized “Oyster garden,” which suggests the two species co‐existed (Miller and Morrison 1988). Presently, Olympia oysters were found in low densities within the eelgrass beds of San Diego Bay (Reed and Hovel 2006). Proximity to eelgrass beds may be advantageous for the oysters, as eelgrass stabilizes sediment and thereby prevents oysters from being smothered (Harris 2004). Although not many reefs structures exist today for

7 use as a reference system, qualitative information can be used to determine the relationship between Olympia oyster reef aggregates found within or near eelgrass.

Ecology of Zostera marina Knowledge of the broader impacts of native oyster re‐establishment on the ecosystem will assist in the restoration process. In particular, understanding the interactions between the native oyster and native eelgrass will aid in the determination of how and where to initiate restoration projects within the Pacific Northwest (PNW). Most estuaries in the PNW have large intertidal areas covered by dense eelgrass beds or existing beds of cultured C. gigas, which is also the same intertidal habitat where Olympia oysters could potentially be re‐established. Therefore, there is a need to understand the relationship between oysters and eelgrass and assess the impacts of restoration efforts.

Distribution Known as common eelgrass, or seawrack, Zostera marina (L., 1753) is the most widely distributed of all seagrasses in the world (Kuo and den Hartog 2001). On the PNW coast, Z. marina can be found from southern British Columbia, Canada, to Humboldt Bay, California. Eelgrass is a marine flowering monocot with rhizomatous root structures and erect, blade‐like leaves. Z. marina grows primarily on mud or mixed mud and sand from +1.5m to ‐1.0 m below MLLW to a depth where they are light‐limited (Bayer 1979, Philips 1984, Hemminga and Duarte 2000). Major stressors include temperature fluctuations below ‐6°C or above +40.5°C; water flows outside of the optimum range of 0.6‐0.8 knots; and salinities < 10 or > 32 (Philips 1984). In the PNW, Z. marina occupies a lower tidal range than its invasive counterpart, Z. japonica (Larned 2003), although the species overlap to some degree (Wisehart 2006, pers. obs.).

Temperate eelgrass systems are seasonally dynamic with respect to growth (Duarte 1989); in Netarts Bay, new growth begins in February, major growth occurs in July, and summer leaves begin to desiccate in November (Kentula 1983). Z. marina produces flowering shoots which are rounded and contain seeds. Flowering stalks are developed from March and through July, seeds germinate from April through July, and seeds are released in mid‐August through October (Kentula 1983). Eelgrass beds are characterized by having more perennial shoots than annual shoots (Kentula and McIntire 1986). Within Netarts Bay, Z. marina exhibits distinct summer and winter morphology; leaves tend to be longer and wider in summer (McIntire 1983).

The long, wide summer leaves are sloughed off as shorter, narrower winter leaves grow (Kentula and McIntire 1986).

Eelgrass and essential fish habitat Eelgrass is an important component of estuarine systems and provides numerous ecosystem services (Costanza et al. 1997, Hemminga and Duarte 2000). Ecosystem services are defined as “The conditions and processes through which natural ecosystems, and the species that make them up, sustain and fulfill human life” (Daily 1997). Ecosystem services provided by eelgrass include sediment stabilization, food for migratory birds (Brant geese), physical structure, nursery grounds for commercially and recreationally important species (Roni et al. 1998), contribution to detrital food web, filtration, and stabilization (Stout 1976, Short and Coles 2001, Orth et al. 2006). Alterations of eelgrass abundance or distribution may affect the capacity of the eelgrass to provide these ecological services (Orth et al. 2006). For instance, if eelgrass declined, its ability to provide habitat would also decline. Eelgrass is considered an indicator species and provides a measure of ecosystem health for a specific locale over time (Thom et al. 2003, Orth et al. 2006).

Eelgrass beds in Oregon are currently managed as essential fish habitat (EFH), defined as “Waters and substrate necessary to fish for spawning, breeding, feeding, or growth to maturity” (PFMC 2006). The concept of EFH was established by the Magnuson‐Stevens Fisheries Conservation and Management Act (Public Law 94‐265). In the Pacific Northwest, EFH management is regulated by the Pacific Fisheries Management Council (PFMC) (2006). The PFMC has management plans for commercial and recreational west coast fisheries. A component of the management plans is the identification of Habitat Areas of Particular Concern (HAPC), which includes eelgrass beds. Certain fisheries in Oregon are threatened and in order to protect these species, alteration of any HAPC is discouraged. Thus, eelgrass management in the PNW is centered on a no‐net loss policy. In Oregon, EFH management is regulated and enforced by the Department of State Lands. This management policy presents restoration practitioners with a challenge given that most oyster restoration projects will take place in intertidal areas that likely contain eelgrass. Therefore, research to define the interactions between oysters and eelgrass is needed to aid in the management of estuarine habitats as EFH. Potentially, Olympia oysters could provide EFH as well (Ruesink et al. 2005), which may alter the definition and management of EFH in the PNW.

Restoration methods Restoration of self‐sustaining populations of the native oyster is conceivable, as the species currently does not face all of the same threats attributed to its decline. First, the current presence of native oysters along the west coast indicates environmental conditions are still favorable for their growth and reproduction. Second, oyster harvests are now regulated and the potential for legal over‐harvest no longer exists. Third, water quality conditions have improved significantly since the passage of the Clean Water Act (33 U.S.C. 1251), which regulates waste dumping. Interest in O. conchaphila restoration resulted in pilot studies and restoration projects in major estuaries such as Tijuana Lagoon (CA), San Francisco Bay (CA), Yaquina Bay (OR), and Puget Sound (WA), among others. A common goal of these projects is to learn about the Olympia oyster and assist in the understanding, development, and implementation of restoration procedures (NOAA Restoration Center 2007).

The goal of the restoration effort described here was to address two potentially limiting factors for populations of O. conchaphila in Netarts Bay: inadequate broodstock and limited settlement substrate (Harris 2004, Brumbaugh et al. 2006). TNC chose to supplement the native oyster population by adding shell cultch set with Olympia oysters to Netarts Bay. This presented the unique opportunity to not only investigate if, and how well, recently transplanted oysters survived, grew, and reproduced, but also examine potential impacts on the abundance and density of the eelgrass bed. To examine these questions, we outplanted cultch in low, medium, and high densities on the eelgrass bed and determined how the addition of shell cultch impacted the eelgrass bed by comparing eelgrass control plots to cultch treatment plots. The objective of the following chapter is to present the analyses of the first year of oyster growth and reproduction after outplanting, along with the project’s initial effects on the eelgrass at the project site, in a restoration ecology context.

Chapter 2: Ostrea conchaphila reestablishment

Introduction The restoration and re‐establishment of native species and habitats can be guided by the principles behind ecological restoration as a discipline. Ecological restoration is defined as “The process of assisting the recovery of an ecosystem that has been degraded, damaged, or destroyed” (Society for Ecological Restoration International (SERI) 2004). All concepts related to restoration are embodied in the term “Ecological restoration,” which includes planning, implementation, monitoring, theory, policy, and more(Temperton and Hobbs 2004). “Restoration ecology” solely represents the theoretical and empirical components of ecosystem development (Temperton and Hobbs 2004). Adaptive restoration is the application of restoration ecology: the utilization of monitoring data to inform management and restoration decisions (Zedler and Callaway 2003). Many projects fall between restoration ecology and adaptive restoration, where ecological data is used in the development of future experimental projects at the restoration site

Ecological restoration can be beneficial, as it often results in increased ecosystem function, complexity, and structure (Ehrenfeld and Toth 1997, Palmer et al. 1997). Identification of specific ecological criteria will aid the development of a guiding image for the restored systems. One method of assessing the progress of a restoration is to examine the ecological trajectory of the system. Trajectories allow us to assess the development of a system and evaluate the restoration, but cannot predict the restoration endpoint. As a system develops along a trajectory from a degraded system toward a restored system, ecosystem function and structure increase according to the ecosystem perspective theory (Ehrenfeld and Toth 1997). Alternatively, as time increases, the ecosystem complexity and function increase as the system reaches a restored state, termed the community perspective theory (Palmer et al. 1997). With an increase in ecological complexity comes an increase in the ecological services provided by the system. Ecosystem services are defined as “The conditions and processes through which natural ecosystems, and the species that make them up, sustain and fulfill human life” (Daily 1997). Common examples include air and water purification, pollination, and flood and drought mitigation. Within the community perspective of ecological trajectories, a degraded system with low ecological complexity and/or function may be missing a component and ecological services

11 may be reduced or absent. Using this same perspective, an intact or restored ecosystem would be capable of performing more ecological services. The restoration will be considered complete when no additional management or restoration is required (SERI 2004).

Native shellfish populations have been highly degraded along the west coast of North America (Kirby 2004) and there is increased interest in Olympia oyster restoration as a culturally and ecologically important species (Peter‐Contesse and Peabody 2005, Dethier 2006). O. conchaphila (= O. lurida, Carpenter, 1857) once was found in west coast estuaries from Sitka, Alaska, to Baja California (Baker 1995). Olympia oysters were extremely popular in the late 19th century and harvests were important to the region’s developing economy (Edmondson 1923, Steele 1957). However, oyster populations began to decline by 1900 due to a combination of overharvest, degraded water quality, and habitat loss due to shell substrate removal (Couch and Hassler 1989, Baker 1995, Kirby 2004, Ruesink et al. 2005). Today the non‐native Pacific oyster, Crassostrea gigas is grown in west coast estuaries for commercial purposes, but it is uncertain whether it replaced the ecological functions of O. conchaphila. C. gigas may occupy different region within estuaries and fill a different ecological role than the native oyster (Ruesink et al. 2005). C. gigas do not form reefs and are harvested every 2‐3 years, and therefore provide short‐term, non‐permanent habitat. Restoration of the native oyster may restore lost ecological functions unique to O. conchaphila. However, in order to re‐establish the native oyster, two currently limiting factors for populations of O. conchaphila must be addressed: inadequate settlement substrate and lack of adequate broodstock (Baker et al. 1999, Harris 2004, Brumbaugh et al. 2006).

Benefits of native oyster restoration include re‐establishment of the ecosystem services (e.g. Daily 1997, Higgs 1997, Menninger and Palmer 2006) provided by the oysters and their reefs (reviewed by Coen and Luckenbach 2000). Oysters are an important foundation species within estuarine systems (Bruno and Bertness 2001, Brumbaugh et al. 2006). Oysters provide habitat for themselves by the creation of shell substrate favorable for spat settlement. Oyster reef assemblages also may function as ecosystem engineers (Jones et al. 1994, 1997) by providing structure for other biota in estuaries (Coen et al. 1999, Gutiérrez et al. 2003, Ruesink et al. 2005). Oyster populations are capable of filtering high water volumes and can consume large amounts of phytoplankton, which may contribute to improved water quality in eutrophied

12 estuaries (Newell 1988). Oyster restoration and reef development may increase biodiversity, from microbes (Nocker et al. 2004) to invertebrates and fishes (Harding and Mann 2001, Tolley and Volety 2005). Species abundances of juvenile and adult fish may increase in habitats in where oyster reefs are situated near seagrass beds (Grabowski et al. 2005). Estuaries which no longer have lost native oysters may have also lost some degree of ecological function; therefore, oyster restoration is an opportunity to regain these functions.

A re‐establishment project was developed to test restoration methods for native Olympia oysters within their historic habitat. Initiated by The Nature Conservancy (TNC) of Oregon, the project represents an interagency Olympia oyster restoration effort in Netarts Bay, Oregon. Although there are few existing reefs for use as a reference system, information on historical reefs guided the restoration effort (Fitch 1953, Steele 1957, Marriage 1958, Espy 1977, Couch and Hassler 1989, Gordon et al. 2001). For instance, historic maps from Yaquina Bay illustrated Olympia oysters as intertidal reefs, bordered by deep channels (Fasten 1931). The long‐term project goal is to re‐establish an aggregate reef system similar to what may have previously existed. TNC addressed previously determined limiting factors to oyster recovery and chose to supplement the native oyster population through the addition of “cultch” to Netarts Bay: a combination of clean Crassostrea gigas substrate shells and newly settled O. conchaphila. Three densities of cultch were outplanted (4%, 11%, and 19% cover).

This re‐establishment project presented a unique opportunity to investigate not only the effectiveness of substrate and broodstock supplementation but also the effects of the restoration on the broader estuarine ecosystem. PNW estuaries are characterized by large intertidal areas with dense eelgrass beds (Zostera marina) (Philips 1984) and have historically included Olympia oysters (Miller and Morrison 1988). Restoration sites within the PNW may be limited to areas not used for navigation or commercial aquaculture, which may contain eelgrass beds. Eelgrass is managed as Essential Fish Habitat (EFH) in the PNW, as it provides many ecological services beneficial to commercial and recreational fisheries (Pacific Fishery Management Council (PFMC) 2006).

The project aimed to identify a threshold cultch density where oysters, cultch, and eelgrass can coexist without detriment to the eelgrass populations. If oyster re‐establishment negatively impacts EFH, future re‐establishment projects may be discouraged. Multiple abiotic

13 and biotic factors and interactions may influence the re‐establishment of the oysters and the dynamics of eelgrass beds at the site (Temperton and Hobbs 2004). This includes tidal inundation patterns, or indirect effects between species (see review by Wootton 1994, Mitsch and Erik 2003). For instance, burrowing shrimp may compete with oysters as filter feeders (Feldman et al. 2000) or macroalgae abundance may be detrimental to eelgrass beds (Hauxwell et al. 2001). The first objective of this study was to determine if the oysters have the capacity to become a self sustaining population through examination of their survival, growth, and reproduction. The second objective of this study was to characterize the re‐establishment site, quantify the effect of oyster density on the abundance of eelgrass, and identify a cultch density that minimizes negative impacts on eelgrass.

Methods

Site description The re‐establishment site is both ecologically ideal and practical for restoration and monitoring purposes. The site is located in one of the few areas of the bay with low navigation traffic, no commercial oyster aquaculture, and no recreational clamming. Netarts Bay (45°26′0″N 123°56′24″W) (Figure 1) has the largest eelgrass beds in Oregon (Stout 1976) and also contains the Oregon Department of Fish and Wildlife’s shellfish research reserve. Two brood years of cultch (Figure 2) were outplanted in four locations within the shellfish reserve in an extensive, relatively uniform eelgrass bed at a 0.0 to ‐1.0 foot tidal height (relative to MLLW) (Figure 3). The first cohort, set in spring 2005 and hereafter referred to as the 2005 brood year, was outplanted in non‐randomized, replicated treatment plots of 0‐4‐11‐19% mean cultch density in locations A and B. The second cohort, set in spring 2006 and hereafter referred to as the 2006 brood year, was outplanted in two additional locations, D and E, located 10 m southwest of location B. Location D contains one continuous plot of 10% cultch while location E contains 6 plots of 14% cultch each. All locations were used in determining oyster growth and reproduction, while only locations A and B were used to evaluate impacts to Z. marina.

Figure 1. Netarts Bay, Oregon, USA. Oregon and Washington emphasized on USA map. Larger inset of outline of Netarts area, with the Bay shown in light gray. Re‐establishment area located within white box. Map adapted from the Oregon Coastal Atlas (Oregon Department of Land et al. 2000).

Figure 2. Shell cultch comprised of O. conchaphila spat on C. gigas shells. Photo of cultch set in May 2007, taken in July 2007.

Figure 3. Map of experimental locations. Relative location of A and B, which are separated by a tidal channel. Cultch treatments are designated by “L” low, “M” medium, and “H” high. Locations D and E are not shown on map. Not to scale. Table indicates total location area, plot size within respective location, brood year, and cultch treatment density. 0‐4‐11‐19% correspond to control, low (L), medium (M), and high (H) treatments, respectively.

L M H Brood Location Total area Plot size year Cultch m‐2 A 1296 m2 122 m2 2005 0‐4‐11‐19% B 1098 m2 144 m2 2005 0‐4‐11‐19% D* 1250 m2 156 m2** 2006 10% L M H E* 864 m2 144 m2 2006 14% *Not shown on map **Subplot size

L M H Location A Location B L L L

LEGEND N L M H M M M channel 4% Shell 11 % Shell 19% Shell

Eelgrass Eelgrass Tidal Control Buffer H H H

Oyster size & growth Oyster shell length (from the base of the umbo to the ventral shell margin) was measured during 2007 at all locations (Table 1, Figure 3). A minimum of 50 O. conchaphila, found by haphazardly tossed 0.25 m2 quadrats, were measured in each plot. Oysters exhibiting morphological characteristics of O. conchaphila with shell lengths <65 mm were considered O. conchaphila. An arbitrary cap of 65 mm was established, as Baker (1995) notes O. conchaphila are rarely >50 mm, therefore oysters >65 mm were excluded from analysis. Oysters >65 mm were morphologically identified as Crassostrea gigas (Kozloff 1996) although it is uncertain how they arrived at the site (Figure 4).

Table 1. Netarts Bay sampling regime, Summer 2007. For each parameter measured (top row), months when data were collected are presented along with the locations where the parameters were measured. Eelgrass measures include percent leaf cover, shoot density, flowering shoot proportion, blade length, and blade width. Blade width was not measured in April. ANOVAs do not include August.

Oyster Oyster Oyster Oyster Eelgrass Density Growth Reproduction Recruitment Measures Month April‐July April‐July May‐August May‐Sept. April‐August Location A, B A, B, D, E A, B, D, E A, B, D, E A, B

Figure 4. Photo comparison of O. conchaphila to C. gigas. One C. gigas is in the center with attached macroalgae, surrounded by several O. conchaphila.

C. gigas

O. conchaphila

Oyster reproduction A subsample of oysters was examined for the presence/absence of reproductive tissue every 4‐6 weeks from May through August (n = 6 events) (Table 1). For each sampling event, 1‐2 pieces of cultch with live O. conchaphila were taken from each medium or high cultch treatment plot (locations A, B, D, E). Gonadal tissue from individual O. conchaphila (n ≥ 20 oysters per event) was viewed under a dissecting microscope to determine if sperm, eggs, and/or larvae were present or absent (total n = 120).

Settlement plates were placed near the treatment locations and surveyed the site for the presence or absence of recruits. For the purposes of this study, recruits are defined as newly settled oysters. Unglazed ceramic tiles were mounted vertically on PVC posts and placed 10‐15 cm above the eelgrass bed surface, and deployed for 2‐4 week intervals (n = 5 events). Shell strings were deployed in August, comprised of ten cleaned and sun‐bleached C. gigas shells, and were collected in September.

It is difficult to morphologically distinguish O. conchaphila recruits from C. gigas recruits, however, DNA barcoding advancements for these species can facilitate species identification (Wang and Guo 2008, Wight et al. 2008). This method compares known genetic markers in mitochondrial DNA (controls) to confirm the species of an individual. Recently recruited spat (<10 mm) were collected from C. gigas substrate at locations D and E (n = 15) in Netarts Bay in September 2007 and preserved in 95% ethanol. DNA was extracted from gill and mantle tissue using a DNAeasy Tissue Kit (Qiagen Inc., Valencia, CA) following manufacturer’s recommendations. Two PCR assays were used: a C. gigas assay and an O. conchaphila assay.

Samples were assayed using PCR conditions described in Wight et al (2008), with minor modifications. PCR reactions were performed in 25 µL volumes containing 1 µL of oyster DNA, 5x PCR buffer, 1.5 mM MgCl, 0.16 mM dNTPs, 0.4 µM forward and reverse primers and 0.04 U/µL Taq polymerase on a GeneAmp 9700 thermocycler (ABI, Foster City, CA) with the following cycle parameters: an initial denaturing phase of 94°C for 5 min, followed by 60 cycles of 94°C for 30 s, 60°C for 30 s and 72°C for 30 s, and finished with a final extension phase of 72°C for 7 min. Amplification products were resolved on a 1.5% agarose gel stained with ethidium bromide. Positive (O. conchaphila) and negative (C. gigas) controls were also used.

Oyster density The density of oysters (oysters per 100 cm2 cultch) was measured monthly in locations A and B (Figure 3). The number of oysters was counted in three haphazardly tossed 0.25 m2 quadrats for each plot. Cultch percent cover was estimated in 5% increments (0‐100%) and included all live and dead shell material, cultch shells, O. conchaphila, and C. gigas. Standardized estimates of oysters per area substrate for each 0.25 m2 quadrat were calculated using Eq. 1.

Eq. 1. 100

where = the estimate of oysters,

= number of oysters counted, C = cultch percent cover/100 Site characterization Basic information used to characterize the re‐establishment site was collected in all locations. Continuous temperature loggers were placed within locations A, B, D, and E from March through September (HOBO Tidbit and Pendant loggers, Onset Computer Corporation). Observations (°C) were made in 30 minute intervals. Salinity at the site was measured with a Horiba Water Checker U‐10 hand‐held probe (Horiba International Corp., Irvine, CA). Qualitative observations of tidal inundation patterns were also made at the site. To further assess site differences, we quantified shrimp burrow hole density in locations A and B in July 2007. Shrimp burrow hole densities were counted in treatment and control plots (n = 24) with five haphazardly tossed 0.25 m2 quadrats. Burrow holes of ghost shrimp (Neotrypaea californienesis) were identified by diameter (<5 mm) and a granular, lined burrow hole with a volcano‐shaped opening, while burrow holes of mud shrimp (Upogebia pugettensis) were identified by the presence of a slimy mucous coating, larger diameter, and a flat surface near the opening (Nesbitt and Campbell 2002). We verified species identification based on burrow holes with a shrimp gun with 100% accuracy (n = 5). Macroalgae percent cover was measured with eelgrass counts to determine if abundance patterns were related to cultch treatments. Measurements were taken within the same three 0.25 m2 quadrats used for oyster density (above): macroalgae percent cover was estimated in 5% intervals (0‐100) and averaged to provide a monthly estimate for each plot.

Eelgrass response to cultch treatments Eelgrass percent cover, shoot density, flowering shoot density, blade length, and blade width were monitored using standard techniques (Duarte and Kirkman 2001) in locations A and B. Measurements were taken within the same three 0.25 m2 quadrats used for oyster density (above) and averaged as a monthly estimate for each plot (Figure 3). The following occurred within each 0.25 m2 quadrat: eelgrass percent leaf cover was estimated in 5% intervals (0‐100); the total number of shoots and flowering shoots were counted; and the length and of a single blade representative of 80% of the other blades within the plot width was measured. Eelgrass measurements from August were incomplete and thus excluded from statistical analyses.

Statistical analyses Three‐way fixed model Analysis of Variance (ANOVA) was used to examine variation in oyster size and eelgrass response variables in relation to cultch treatments (α = 0.05, SPSS statistical package ver. 15.0.1.1). Data were examined for normality and homogeneity of variance (Levene’s or the Hartley’s F‐max tests for homogeneity of variance). Outliers were identified by boxplots, graphs, and plots of residuals and Cook’s distance. Transformations of response variables were used to meet assumptions for normal distributions (noted with each parameter). When data met the normality assumption but variances were not homogeneous, we proceeded with the ANOVA, considered robust with large, relatively equal sample sizes, using Type III sum of squares for unequal sample sizes (Quinn and Keough 2002). For oyster, eelgrass, and macroalgae measurements, a total of 24 total plots were measured with three fixed factors: location (A, B), month (varies by parameter), and cultch treatment (control, low, medium, high); and four interaction terms (96 total degrees of freedom) (Table 1, Figure 3). Games‐Howell post‐hoc multiple comparisons were used when variances were heterogeneous (Toothaker 1993, Kirk 1995). If data did not meet the assumption of normality but had homogeneous variances, non parametric statistics were used to determine differences between locations, months, and treatments. The Mann‐Whitney test was used to compare two groups, while the Kruskall‐Wallis Rank Sum procedure was used for more than two groups (SPSS statistical package ver. 15.0.1.1). The relationships among shrimp burrow hole densities and eelgrass factors were quantified using linear least‐squares regression (Microsoft Excel 2007).

Only one of the nine subsamples within the low density treatment was measured in location B in May. Data were generated for these missing eight subsamples by mean value

20 replacement for this particular month, location, and treatment. Valid data were available for the months of April and June, and it was assumed that between these time points, the eelgrass data points for May follow a similar linear trend. A regression line was fit to the monthly means of the location B, low density treatment plots for the following variables: cultch cover, oyster density, eelgrass percent cover, eelgrass shoot count, flowering shoot count, blade length, and blade width. From this regression line, 95% confidence intervals were fitted and eight random numbers were generated from a uniform distribution within the 95% confidence interval surrounding the estimated variable mean for the month of May. For the remaining data, outliers or missing data (failure to record) were not replaced and resulted in additional unequal subsample sizes.

Results

Oyster size and growth Both brood years of Olympia oysters showed increases in size throughout 2007. The mean size of the 2005 brood year increased by 10.5 mm, with major increases throughout June, July, and September 2007 (Figure 5). Size of the 2005 brood year was significantly different among months (ANOVA, p<0.001, Appendix A), with significant differences between June, July, and September (Games‐Howell; p ≤ 0.002). The 2006 brood year steadily increased in size throughout 2007 with a mean increase of 16.2 mm (Figures 5, 6).

The size frequency distributions (Figures 7, 8) provide detail on the progression of the population and allowed detection of subtle changes along with large recruitment events. The size distributions of the 2005 brood year (Figure 7) stayed relatively uniform from March through September although mean size increased. The size distribution for the 2006 brood year (Figure 8) broadened by September, due to the presence of larger individuals. For the 2006 brood year, there was no large recruitment observed but a small number of oysters remained in the 0‐10 mm size class. This may represent continual small scale recruitment to the site.

Figure 5. Oyster monthly mean size in 2007. Oyster monthly mean linear shell length, where shell length is from the umbo to the furthest point on the shell (mm), ± SE, during 2007.

45 (mm)

35 Length

30 Shell

25 Linear

20 Monthly

Mean 10 Mar Apr May Jun Jul Sep

2005 Brood Year 2006 Brood Year

Figure 6. Mean monthly oyster growth by treatment. Oyster monthly mean size ±SE by treatment (low, medium, high) for the 2005 brood year, from March through September, 2007.

40 (mm)

growth

35 oyster

30 Mean

25 Mar Apr May Jun Jul Sep

Low Med High

Figure 7. Size distribution of 2005 brood year. Figure 8. Size distribution of 2006 brood year. Arrow represents monthly median. Size (mm) on x‐ Arrow represents monthly median. Size (mm) on x‐ axis, proportion on y‐axis. axis, proportion on y‐axis.

0.35 March 2007 0.35 March 2007 0.30 0.30

0.25 0.25

0.20 0.20

0.15 0.15

0.10 0.10

0.05 0.05

0.00 0.00 0 102030405060 0 102030405060

0.35 April 2007 0.35 May 2007 0.30 0.30

0.25 0.25

0.20 0.20

0.15 0.15 0.10 0.10 0.05 0.05 0.00 0.00 0 102030405060 0 102030405060

0.35 May 2007 0.35 June 2007 0.3 0.30

0.25 0.25

0.2 0.20 0.15 0.15 0.1 0.10 0.05 0.05 0 0.00 0 102030405060 0 102030405060

0.35 June 2007 0.35 July 2007 0.3 0.30 0.25 0.25 0.2 0.20 0.15 0.15 0.1 0.10 0.05 0.05 0 0.00 0 102030405060 0 102030405060

0.35 July 2007 0.35 August 2007 0.3 0.30 0.25 0.25 0.2 0.20 0.15 0.15 0.1 0.10 0.05 0.05 0 0.00 0 102030405060 0 102030405060 0.35 September 2007 0.35 September 2007 0.3 0.30 0.25 0.25 0.2 0.20 0.15 0.15 0.1 0.10 0.05 0.05 0 0 102030405060 0.00 0 102030405060 Linear shell length (mm) Linear shell length (mm) 23

Oyster reproduction Oysters from both brood years in all locations contained reproductive tissue (Table 2). Of the 120 oysters examined, 103 oysters (85.8%) contained reproductively active tissue. Reproductive oysters were either identified as males containing sperm (n = 99) or females brooding larvae (n = 4). No eggs were positively identified. Male shell lengths ranged from 2.14‐ 5.18 mm, while brooding females ranged from 2.80‐3.95 mm. Non‐reproductively active oysters ranged from 1.61‐4.16 mm. Larvae were found in May and June, while the highest number of inactive oysters was observed in July (Appendix B).

Recruitment at the site was not detected on any of the ceramic settlement plates or shell strings placed at the re‐establishment site. However, recruits were observed at the site on cultch within the treatment plots. A slight increase of small oysters (< 10 mm) was indicated by the September size distribution (Figure 8). These oysters could be new recruits or possibly slow‐ growing oysters. Additionally, routine field observations detected a settlement event in late September (Figure 9), when oysters < 5 mm were discovered on PVC plot markers and on cultch bags containing a brood settled in May 2007 (not measured in this experiment).

DNA analysis confirmed the existence of O. conchaphila recruits at the site. The O. conchaphila assay shows that PCR products for each of the new recruit samples were of expected size (~150 bp) and were consistent with positive controls for O. conchaphila (Figure 10). The C. gigas assay shows that only the positive control produced a band (Figure 11). An additional O. conchaphila recruit collection and identification assay was attempted in January 2007, but no surviving recruits could be located.

Table 2. Oyster reproductive tissue. Oyster reproductive tissue found by brood year and location. Totals of male, larvae, and non‐active oysters are provided. No females were found.

Reproductive Tissue Type Brood Year Location Male Larvae Non‐active Total 2005 A 34 1 4 39 2005 B 28 1 4 33 2006 C 19 2 4 25 2006 D 18 0 5 23

Total99417120

Figure 9. Photo of September recruitment event. Recruits visible in white circle on C. gigas shells within bags.

Figure 10. PCR assay for O. conchaphila. O. conchaphila PCR assay shows samples corresponding with O. conchaphila positive control. Lane assignment (row 1): ladder (L), O. conchaphila samples 1‐8, C. gigas negative control (NC); O. conchaphila positive control (PO); C. gigas positive control (PC), L. Lane assignment (row 2): L, O. conchaphila samples 9‐15, NC, PO, PC, L.

Row 1:

Row 2:

Figure 11. PCR assay for C. gigas. C. gigas PCR assay shows only the positive control (PC) for C. gigas producing a band. Lane assignment (row 1): ladder (L), O. conchaphila samples 1‐8, negative control (NC), C. gigas positive control (PC), O. conchaphila positive control (PO). Lane assignment (row 2): O. conchaphila samples 9‐15, PC, NC, PO, L.

Row 1:

Row 2:

Oyster density The oyster‐eelgrass interaction plots within locations A and B contained three treatments of cultch density which had different oyster densities. There was a non‐significant positive trend of standardized oyster density (O. conchaphila 100 cm‐2cultch) as cultch percent cover increased (Figure 16). Standardized oyster densities showed no significant interaction terms or factors of location or month, but cultch treatments were significantly different (p < 0.001, log‐transformed ANOVA, Appendix C). The standardized density of oysters was significantly less in the low treatment plots but there was no significant difference between oyster densities in the medium and high treatment plots (Games‐Howell post‐hoc comparisons). Throughout the oyster surveys, C. gigas was found growing on the cultch, which were considered additional substrate included in the shell areal estimates.

Figure 12. Olympia oyster density as compared to cultch treatment. Mean Olympia oyster density (O. conchaphila 100 cm‐2 cultch) by cultch treatment ± SE. “Low” refers to 4% cultch cover, “Medium” to 11% cultch cover, and “High” to 19% cultch cover. Significance at 95% confidence level indicated by (*).

6 5.09

5 shell 3.71 2 ‐ 4 cm

100 3 * 1.85 oysters 2

Olympia 1

0 "Low" "Medium" "High" Cultch treatment

Site characterization Daily mean mudflat surface temperatures ranged from 9.9 (March 27) to 21.3°C (July 4) at the site (Appendix D). Salinities at the site ranged from 26.3 (May) to 33.7 (June) (Appendix E). Qualitative observations indicated that each location had different water flow patterns depending on tidal stage. Location A was generally exposed 30‐45 minutes before location B. Parts of location B were rarely exposed, and locations D and E often retained ≥ 10 cm of water at low tide.

There were measurable differences between the eelgrass beds of location A and location B. Eelgrass within control plots in location A had lower percent cover, higher shoot density, fewer flowering shoots, shorter blades, and narrower blades than eelgrass control plots within location B (Table 3). Seasonal shifts in mean eelgrass percent cover and mean shoot density were evident in both locations, although location A was more variable than location B (Figures 12, 13, 14). Month was consistently a significant factor in the ANOVA analyses for the parameters of eelgrass percent cover, shoot density, flowering shoot percentage, and blade length (see pages 31‐32, Appendix F).

Table 3. Eelgrass dynamics in control plots for locations A and B. Means for eelgrass percent cover, shoot density, flowering shoot percentage, blade length, and blade width within control plots of locations A and B are presented ± SE.

Parameter Location ALocation B Percent cover 73.8% cover ± 4.8 SE 88.6% cover ± 1.7 SE Shoot Density 693 shoots m‐2 ± 86 SE 464 shoots m‐2 ± 21 SE Flowering Shoot %6.3 % ± 2.9 SE 7.0 % ± 4.9 SE Blade Length 36.1 cm ± 10.2 SE 60.29 cm ± 14.2 SE Blade Width 5.17 mm ± 6.1 SE 6.81 mm ± 9.5 SE

Figure 13. Mean eelgrass percent cover. Monthly mean eelgrass percent cover to nearest 5%. Location A designated by (a) and location B by (b). Cultch treatments designated by lines. Means ± SE from August and September are in figures but not in ANOVA analyses.

100 (a) Location A 100 (b) Location B

80 80 cover cover

leaf leaf

60 60 percent percent

40 40 Eelgrass Eelgrass

20 20 Apr May Jun Jul Aug Sep Apr May Jun Jul Aug Sep Control Low Medium High Control Low Medium High

Figure 14. Mean eelgrass shoot density. Eelgrass shoots m‐2 by location ± SE. Shoot counts are reported m‐2. Means and standard errors from August are included in figures but were not included in the ANOVA analyses. Note y‐axis starts at 200.

1200 (a) Location A 1200 (b) Location B

1000 1000 2 2 ‐ ‐ m m

800 800 count count

shoot shoot

600 600 Eelgrass Eelgrass 400 400

200 200 Apr May Jun Jul Aug Apr May Jun Jul Aug Control Low Medium High Control Low Medium High

Figure 15. Mean proportion eelgrass flowering shoots of total shoots. Shown by location ±SE. The peak of flowering shoots was in June, where flowering shoots accounted for 9% of the total shoots.

14 (a) Location A 14 (b) Location B

12 12 2 2 ‐ ‐ m m

10 10 shoots shoots 8 8

6 6 flowering flowering

4 4 Percent Percent 2 2

0 0 Apr May Jun Jul Apr May Jun Jul

Control Low Medium High Control Low Medium High

Burrowing shrimp populations were different between Locations A and B. Ghost shrimp (N. californienesis) and mud shrimp (U. pugettensis) were present in Locations A and B. Mean ghost shrimp burrow densities were similar in both locations (mean = 2.11, stdev = 3.74, median = 4, mode = 0) (Kruskall‐Wallis, p > 0.05). Mean ghost shrimp burrow densities were greatest in high cultch treatments, but non‐parametric tests indicated there were only marginally significant differences between treatments (Kruskall‐Wallis, p = 0.048). In contrast, mud shrimp burrow hole densities were significantly greater in location A than location B (Mann‐Whitney, p < 0.001) (Figure 16). The assumption of homogeneity of variance was not met because of the extreme difference in burrow means by location; therefore non‐parametric tests were used to analyze each location separately. Within location A, there were significant differences of mud shrimp burrow density among treatments (Kruskall‐Wallis p = 0.002), but there was no sequential relationship between mud shrimp burrow density and cultch treatment (overall rank: low < high< control < medium). There were significant differences of mud shrimp burrow density between treatments in location B (Kruskall‐Wallis, p = 0.007), but similar to location A, there was no relation to increasing cultch density (rank: low < medium < control < high).

Figure 16. U. pugettensis burrow hole density. Mud shrimp burrow holes m‐2 for location A (dark bars, left axis) and location B (light bars, right axis) with separate scaling shown with standard error. Mean burrow density for location A = 98.4 ± 6.70 SE burrow holes m‐2. Mean burrow density for location B = 4.07 ± 0.85 SE burrow holes m‐2.

160 Location A Location B 14 140

12 B A

120 10 Location Location 100

, , 2 2

8 ‐ ‐ m m

80 6 holes

holes 60

4 40 Burrows Burrows 20 2

0 0 Control Low Medium High

Linear regressions were used to investigate relationships between shrimp burrow densities and eelgrass percent cover, and eelgrass shoot density. In location A, no relationships were evident between burrow hole density and eelgrass percent cover (r2 = 0.136, F = 0.313, p = 0.949, df = 2) or between burrow hole density and eelgrass shoot density in location A (r2 = 0.022, F = 0.046, p = 1, df = 2). In location B, a scatterplot suggested a negative linear relationship between burrow density and eelgrass percent cover (r2 = 0.450, F = 1.638, p = 0.452, df = 2), however, there was no relationship between burrow density and eelgrass shoot density in location B (r2 = 0.181, F = 0.441, p = 0.884, df = 2).

Macroalgae (primarily Enteromorpha spp.) was present at the re‐establishment site throughout the summer season and field observations indicated a negative effect on eelgrass populations (Figure 17, Appendix G). Mean abundance peaked in June and macroalgae remained at the site through September. The distributions were non‐normal and heteroscedastic, therefore non‐parametric tests were used and neither location (Mann‐ Whitney, p = 0.255) nor treatment (Kruskall‐Wallis, p = 0.070) were significant factors in the distribution of macroalgae. Macroalgae abundance was significantly different for each month (Kruskall‐Wallis, p<0.001). Patterns of mean distribution suggested a relationship to cultch despite the marginally non‐significant Kruskall‐Wallis test results. An ANOVA on the rank‐ transformed values for June indicated both location and cultch treatment as significant factors affecting macroalgae distribution (Table 4). Games‐Howell multiple comparisons indicated the control treatment was significantly different than the high density cultch treatment (p = 0.008), but there was an increasing trend of macroalgae with increasing cultch density.

Figure 17. Mean macroalgae percent cover by treatment. Mean macroalgae percent cover ±SE for all treatments in locations A and B by month.

30 cover

percent 20

Macroalgae 5

0 Apr May Jun Jul Aug

Cont Low Med High

Table 4. ANOVA on rank‐transformed values of macroalgae percent cover. Type III Sum of Source Squares df Mean Square F Sig. Location 189.951 1 189.951 5.095 0.037 Cultchtrt 487.740 3 162.580 4.361 0.018 Location * Cultchtrt 21.855 3 7.285 0.195 0.898 Error 671.021 18 37.279 Total 6113.000 26

Eelgrass response to cultch treatments The response of eelgrass parameters to cultch treatments was similar for both locations, despite differences in the eelgrass bed dynamics at each location and after accounting for seasonality. Mean eelgrass percent cover and shoot density were often greater in the control and low cultch treatment plots when compared with the medium and high treatment plots (Figures 12 and 13, Appendix H). An ANOVA on eelgrass percent cover yielded no significant interaction terms but the factors of location, cultch treatment, and month were significant contributors to variances in eelgrass percent cover (ANOVA on ArcSin transformed values, Table 4). Due to heteroscedasticity, Games‐Howell multiple comparisons were used to identify differences among cultch treatments. High treatments had significantly less eelgrass cover than

32 controls (p = 0.010), and low treatments were different than medium treatments (p = 0.005) and high treatments (p < 0.001). Eelgrass shoot densities also declined in high treatment plots. The interaction between location and month was significant (Table 5). Eelgrass shoot densities exhibited different patterns of location and month, contributing to the significant interactions terms. However, shoot densities had similar response patterns after other sources of variation were accounted for. Games‐Howell multiple comparisons indicated that control and low treatments had significantly greater shoot densities than the high treatment.

The response of eelgrass flowering shoot percentages to cultch treatments was less clear. The count of flowering shoots in April was essentially zero for all plots. To increase the effectiveness of the analysis, the month of April was removed from the ANOVA performed on the log‐transformed standardized flowering shoot percentages. The location*month and treatment*month interaction terms was significant (Table 6). Differences between cultch treatments were only marginally significant (p = 0.045), and Games Howell multiple comparison procedures report no differences between cultch treatments.

Growth morphology of eelgrass did not show patterns associated with cultch treatments. There were no discernable patterns in eelgrass blade length with the exception of seasonality (Appendix I). An ANOVA on the log‐transformed values found no interaction terms to be significant, but the factors of location, cultch treatment, and month were all significant contributors to variation in blade length (Appendix J). The low treatment had longer blade widths than the high treatment (p = 0.033), but there were no significant differences between the control and any of the treatments (p > 0.05). There were differences in mean eelgrass blade width among cultch treatments, but no linear pattern coincided with cultch treatment (Appendix K). No significant interaction terms were found in an ANOVA on the log‐transformed values of eelgrass blade width. Location (p<0.001) and cultch treatment (p = 0.011) were significant factors, while month was not (Appendix L). There were no differences between cultch treatments under Games‐Howell post‐hoc comparisons.

Table 5. ANOVA of ArcSin transformed values of eelgrass percent cover. Type III Sum of Source Squares df Mean Square F Sig. Location 0.180 1 0.180 6.967 0.011 Month 0.506 3 0.169 6.527 0.001 Cultchtrt 0.904 3 0.301 11.663 <0.001 Location * Month 0.127 3 0.042 1.642 0.189 Location * Cultchtrt 0.196 3 0.065 2.531 0.066 Month * Cultchtrt 0.120 9 0.013 0.516 0.857 Location * Cultchtrt *Month 0.277 9 0.031 1.191 0.318 Error 1.525 59 0.026 Total 112.936 91

Table 6. ANOVA of log transformed values of eelgrass shoot count m‐2. Type III Sum of Source Squares Df Mean Square F Sig. Location 0.753 1 0.753 72.936 <0.001 Month 0.529 3 0.176 17.103 <0.001 Cultchtrt 0.349 3 0.116 11.281 <0.001 Location * Month 0.132 3 0.044 4.252 0.009 Location * Cultchtrt 0.058 3 0.019 1.877 0.143 Month * Cultchtrt 0.056 9 0.006 0.608 0.786 Cultchtrt * Location *Month 0.162 9 0.018 1.743 0.099 Error 0.619 60 0.010 Total 655.398 92

Table 7. ANOVA of log‐transformed values of percent flowering shoots. Type III Sum of Source Squares df Mean Square F Sig. Location 0.117 1 0.117 5.756 0.021 Month 1.601 2 0.801 39.307 <0.001 Cultchtrt 0.178 3 0.059 2.913 0.045 Location* Month 1.095 2 0.548 26.881 <0.001 Location * Cultchtrt 0.085 3 0.028 1.386 0.259 Month * Cultchtrt 0.297 6 0.050 2.433 0.041 Location* Cultchtrt * Month 0.111 6 0.019 0.911 0.496 Error 0.896 44 0.020 Total 46.652 68

Discussion

Reestablishment of oysters within Netarts Bay The process of ecological restoration may take many years, but practitioners often need to make management decisions based on the information available at the time. Monitoring and evaluation provide information on whether or not the system is recovering and regaining its capacity to provide ecosystem goods and services. The overall goal of the project is to re‐ establish a self‐sustaining population of native oysters within Netarts Bay, Oregon. Ideally, a self‐sustaining population of oysters will produce enough broodstock and shell substrate for the population to survive without additional anthropogenic management or additions to the system (Coen and Luckenbach 2000). Once the oyster population is growing, reproducing, recruiting, and those recruits survive to reproduce, it will be considered re‐established.

Our results indicate that Olympia oysters are capable of growth, reproduction, and recruitment in their historic habitat and have the potential to become a self‐sustaining population. In September 2007, the Netarts Bay oysters were in the upper range of reported size values, slightly larger than previously reported sizes of 35‐45 mm in size in 3 years (Baker 1995, Peter‐Contesse and Peabody 2005). The overall mean size of each brood year increased throughout 2007, although the brood years demonstrated different growth patterns. The mean oyster size of the 2005 brood year did not increase until June, while the 2006 brood year exhibited constant increases in mean size from March to September. This may have been due to location or different growing conditions at the time each brood year was set. The early‐season delay in growth for the 2005 brood year may possibly be related to the reallocation of energy by older oysters from growth into reproductive materials (Coe 1932).

The reproductive capacity of the oyster populations provided evidence of restoration progress. O. conchaphila are protandrous (male stage develops first), larviporous (females brood larvae), and hermaphroditic (switches between male and female reproductive forms) (Coe 1932). Coe (1932) reported that spermatogenesis begins within 12 weeks of attachment, and oogenesis begins at 22‐30 weeks. Dissections began at 104 and 52 weeks after settlement (of the 2005 and 2006 brood years, respectively) when reproductively active oysters from both brood years were found. Spermatogenesis was primarily observed, which is typical of young oyster populations (Coe 1932). A limited number of oysters from both brood years contained

35 larvae, but as the population ages, the number of females present and brooding larvae is expected to increase.

Larval recruitment also provided an indication of the oyster population restoration progress. The site is within range of a viable seed source, which indicates the site has enough potential to invest resources in additional restoration efforts. In 2007, two recruitment events apparently occurred by were not evident in the size distributions (Figure 7, Figure 8). Early research on O. conchaphila found multiple spawning events within a year are possible (Coe 1932). One 2007 event may be present at the 10 mm mark in September (Figure 8). The other event was observed (after qualitative monitoring concluded) in late September on cultch in the treatment plots and cultch bags containing a cohort settled in May 2007 (Figure 9). This suggests the possibility that both 2005 and 2006 brood years may also contain two age classes. However, as illustrated by the size frequency distributions and the rapid growth of first year oysters, a second age class might be easily missed because its height values overlap with the cohort set earlier that particular year.

Oyster density and reef structures A critical element of oyster re‐establishment is to identify whether the oysters are able to produce enough shell to build reef aggregates and continue to recruit new cohorts (Powell and Klink 2007). If the Netarts Bay population is not capable of producing its own shell for younger generations of oysters, it will require management through continued substrate addition and/or broodstock supplementation. Within this study, the cover and density of oysters within treatments was measured as an indirect measure of survival. Changes in cultch and/or oyster density may indicate potential changes in growing conditions and provides an estimate of the survival of outplanted oysters. The low treatment plot had the lowest oyster density, while the medium and high treatment plots had greater oyster densities but were not statistically different from each other. This indicates a moderate density of oysters can be achieved by outplanting shells at the medium (11%) density instead of the high (19%) density.

It is unknown how these experimental densities compare to the qualitatively described oyster populations and reef aggregates in Baker and references therein (1995). Continued monitoring of the site will likely provide insight into how oyster reef aggregates form (see recommendations, p 38). Other reef descriptions are rare but are still useful in assessing the

36 progress of the Netarts Bay restoration. In Elkhorn Slough, CA, oysters are found in naturally occurring clumps (< 0.5 m2) or a reef on an anthropogenic rock bed (> 0.5 m2), with a maximum density of 340 oysters/m‐2 (Heiman et al. 2008). At these densities, even the Netarts Bay high density treatment (0.19 m2) is not considered a “reef,” but may resemble the natural “clump.” We are unsure of what the aggregate reef environment once characteristic of O. conchaphila looks like, but perhaps it is a hybrid between the “clumps” and the “reef.”

Interactions between cultch and eelgrass The second objective of the project was to quantify the impacts of the re‐establishment efforts at the restoration site. Before describing the impacts of cultch on eelgrass, it is important to establish that the experimental locations were very different from one another. Z. marina beds are often heterogenous and have high spatial and temporal variability (Duarte 1989, Hemminga and Duarte 2000, Thom et al. 2003). The two locations in this study were very different from each other with respect to their drainage patterns, burrowing shrimp populations, and eelgrass beds. Location A generally had higher shoot counts but an overall lower eelgrass leaf cover than location B, which had low shoot counts and high percent leaf cover. This is consistent with the pattern described by Kentula and McIntire for Netarts Bay (1986). Seasonal changes in eelgrass were evident at the experimental site, where eelgrass percent cover, shoot density, and blade length increased from spring through summer and declined into fall.

Even though eelgrass populations in both locations had high variability and there were significant seasonal influences, cultch treatment remained a statistically significant factor throughout the analyses. High cultch treatments had lower eelgrass percent cover and shoot density. Variability in eelgrass leaf cover, shoot density, and flowering shoot density was attributed to cultch treatment. Both locations experienced declines in eelgrass cover and shoot density with each increasing cultch density treatment. Eelgrass blade length and blade width did not respond uniformly to cultch treatments and, therefore, their use for indicating short‐term changes as a result of cultch addition is not recommended.

Eelgrass at the re‐establishment site had relatively high percent cover estimates and low shoot densities (Figure 13, Figure 14, Appendix H). Netarts Bay eelgrass beds have greater shoot densities than other PNW estuaries (see Thom et al. 2003). An in‐depth study of the Netarts Bay

37 eelgrass beds was conducted at a higher tidal elevation (+1.0 to +1.4 feet) using smaller (0.04 m2) quadrats and reported double, or even triple, the shoot density reported in this study (Kentula 1983, Kentula and McIntire 1986). A potential difference in the eelgrass patterns may be that the TNC restoration site is at a lower tidal height than the previous study. Eelgrass found at higher elevations (+1.0 feet) is usually the annual growth form, while eelgrass found at lower tidal elevations (‐1.0 feet) is most likely the perennial growth form (Bayer 1979). This oyster re‐ establishment site is at lower tidal elevation (0 to ‐1.0 feet) and most likely contains perennial eelgrass.

Additional ecological interactions An unforeseen impact at the re‐establishment site was the appearance of large clumps of macroalgae. Macroalgae is known to have a negative impact on seagrass beds, causing damage and/or die offs similar to those observed at the site (Valiela et al. 1997, Raffaellii et al. 1998). Enteromorpha spp. became anchored to eelgrass, cultch, and plot markers within the sites. Eventually, these clumps caused small (< 3 m2) eelgrass die‐offs as the season progressed due to smothering and anoxic conditions. This provides evidence that, in addition to oyster cultch, other ecological interactions may be impacting eelgrass abundance and growth patterns. The presence of cultch may abet attachment of Enteromorpha spp. and cause an increased macroalgal load to the area. However, it is important to recognize the decline in eelgrass percent cover and shoot density within the high cultch density was apparent in April and May (Figures 12 and 13), before the macroalgae bloom in June. As a result, oyster re‐establishment activities may indirectly facilitate the degradation of eelgrass beds but additional long‐term research is needed.

The mud shrimp burrow hole densities were much greater in location A than location B. However, within location A, no relationship was found between the burrow density and eelgrass percent cover or the eelgrass shoot density. This evidence indicates eelgrass patterns were unrelated to shrimp density and the shrimp densities likely did not confound the cultch treatment results. U. pugettensis densities at Netarts Bay location A were similar to those estimated for Willapa Bay, WA (Dumbauld et al. 1996). Compared to Oregon estuaries, the Netarts Bay location A densities were relatively low (Thom et al. 2003, DeWitt et al. 2004). In

Coos Bay, OR, eelgrass areas exceeding 100 shoots m‐2 had low burrow densities (<5 m‐2) (Thom et al. 2003).

Recommendations Detailed understanding of reproductive cycles and recruitment patterns of the Olympia oyster will enhance restoration efforts within Netarts Bay. As the site is within range of a seed source, broodstock may no longer be a limiting factor. If so, re‐establishment efforts could focus on shell substrate supplementation. If additional cultch outplantings are to occur, it may be beneficial for managers to plan for “recruitment enhancement” (sensu Mann 2007), coordinating the outplanting with a bay‐wide recruitment event to increase settlement. Reproductive monitoring should start before oogenesis begins, typically when water temperature reaches 13oC ‐ 16oC for 3‐6 months (Baker 1995 and references therein). Along with water column sampling, oyster dissections can aid in the detection of spawning events. After population densities increase and can sustain weekly harvests, weekly monitoring of reproductive status should occur in March before the water reaches 13oC, as reproductive requirements may have changed. However, in order to avoid destroying potential broodstock in a small population, reproductive sampling may be limited to small sample sizes. Attempts to capture recruitment should be made consistently on a weekly basis throughout the reproductive season. To obtain measures of recruitment, materials such as shell cultch and PVC, which the Olympia oysters prefer to set upon, should be used. Monitoring the survival of outplanted oysters and current and future recruits will provide additional information on status of the re‐ establishment. This study provided qualitative descriptions of settlement events in 2007, but quantitative monitoring of the survival of individual oysters to reproductive age will indicate re‐ establishment progress and the point when broodstock supplementation is no longer needed.

Although native oyster restoration on the west coast has only recently gained momentum, large scale oyster restoration on the east coast has been underway for decades. Some of the same recommendations for C. virginica restoration efforts (Coen and Luckenbach 2000) apply to the restoration and re‐establishment of O. conchaphila. Other recommendations include predicting recruitment, investigating disease events, quantifying biofouling of cultch substrate, and quantifying predation rates of both native and non‐native species (Mann 2007). Netarts Bay observations indicated that the ability of larvae to settle and survive may be

39 affected by biofouling of the substrate cultch by algae or bryozoans and predation by native and non‐native predators including Carcinus maenas. Another challenge to oyster recruitment and survival may be related to certain types of bacteria, i.e. Vibrio spp., which are harmful to oysters in their early life forms and are common in the Pacific Northwest (Estes et al. 2004). Vibrio spp. has the capacity to kill larval C. gigas and O. edulis in the water column (Estes et al. 2004). Netarts Bay is home to extensive C. gigas aquaculture along with the Whiskey Creek Shellfish hatchery, and Vibrio tubiashii outbreaks have occurred within the hatchery within the last few years (M. Wittwer, pers. comm.). These hatchery bacterial outbreak events may have an impact on juvenile O. conchaphila as well, which may impact the ability of the Olympia oysters to become a re‐established and self‐sustaining population.

Future research Along with continued monitoring of oyster density, future research may focus on qualifying and quantifying both the direct and indirect effects of the oyster re‐establishment. One particular effect is the presence of C. gigas at the re‐establishment site. Several oysters larger than 65mm were found at the site, which morphologically looked like C. gigas and not O. conchaphila. These oysters were presumed to be C. gigas and were included in the “cultch substrate” estimates, as O. conchaphila is known to settle on commercially grown C. gigas. The source of the C. gigas is unknown, although we presume the C. gigas cohorts were set in the same years as the O. conchaphila (2005 and 2006, respectively). The C. gigas compete with O. conchaphila for space on the shell substrate, and may crowd or outgrow them (per. obs.). It is unknown whether C. gigas and O. conchaphila compete for food sources, although qualitative evidence indicates O. conchaphila may be at a disadvantage (Elsey 1935). A quantitative feeding study will provide more conclusive evidence. If the oysters directly compete for resources, this may affect restoration planning, as certain areas of the bay may contain different quantity and quality phytoplankton (Newton and Horner 2003).

As oyster restoration projects are developed, it remains critical to address any potential impacts the oysters and subsequent cultch will have on the eelgrass beds. As aggregate reefs form within eelgrass beds, there may be changes to both oyster densities and eelgrass abundances. These results indicated a negative impact of high (19%) density cultch on eelgrass percent cover and shoot density. To confirm this effect, a higher cultch density, between 20 ‐

50% should be tested. Typically, eelgrass densities are lower in areas of higher oyster densities (Tallis et al. 2007). Determining the interactions between O. conchaphila and native eelgrass will also facilitate the understanding and management of EFH. Because of the current no‐net loss policy, restoration activities which may negatively impact eelgrass abundances could be discouraged. However, it is possible that Olympia oysters and eventually reef structures may be incorporated into the definition of EFH. Other changes within the eelgrass bed may occur as a result of the restoration and need to be studied as well. For instance, prior to summer 2007, there was an existing bed of Z. marina and Z. japonica south of location B, but as summer 2007 progressed, Z. japonica shoots began to appear within location B. In addition, macroalgae presence at the site also has a strong impact on eelgrass, as previously noted by Kentula and McIntire (1986). If the presence of shell cultch facilitates the attachment of macroalgae such as Enteromorpha spp., this may have consequences for both oysters and eelgrass due to the anoxic conditions created by decaying macroalgae (den Hartog 1994). The impacts of the oyster re‐ establishment will be understood with time, which illustrates the importance of long‐term eelgrass monitoring at the site.

Conclusions This project is the first step towards increasing the understanding of the process of oyster re‐establishment, reef formation, and potential impacts of oysters on eelgrass. Through the intensive monitoring effort during 2007, it is clear the Netarts Bay oysters are growing, reproducing, and recruiting to the site. Both brood years are increasing in size and were reproductively active; some oysters were found brooding larvae. The medium cultch treatment was identified as a potential threshold where oysters exist at a high density without statistically significant negative cultch impacts to eelgrass percent cover or shoot density. The long‐term impacts to the eelgrass beds are unknown. Although the limiting factors of broodstock and shell substrate to oyster recovery are being addressed, it became apparent that several other ecological interactions may inhibit or delay the re‐establishment of O. conchaphila. To achieve a self‐sustaining population, these external factors must also be addressed (Palmer et al. 2005).

As oyster restoration projects increase in the PNW, it is likely that available project sites will contain eelgrass and there is a need to understand the relationship between oysters and eelgrass. Because Netarts Bay is a conservation estuary designated for limited development but

41 continued resource consumption, a compromise must be reached between complete ecosystem restoration and anthropogenic needs such as aquaculture or land development (Weinstein 2007, 2008). This is supported by the beliefs of PNW residents who want a balance between environmental protection and economic considerations (Huppert et al. 2003). Future restoration efforts within the PNW may benefit from plans to develop a long‐term Olympia oyster restoration vision to guide projects and efficiently utilize resources, guided by public input (Palmer et al. 1997, Palmer et al. 2005). This vision should include a clear definition of restoration evaluation criteria and the ways and means to apply such criteria to the project (see Short et al. 2000). Adaptive restoration through community support, practical guidance, and scientific testing will be a critical component of the re‐establishment of the native oyster to the west coast.

Implications for practice

• The 11% (medium) cultch treatment had the highest density of oysters without statistically significant impacts on eelgrass abundance. • High cultch densities had statistically significant, short‐term, negative impacts on eelgrass abundance. • Eelgrass percent cover and eelgrass shoot density were greatest in control plots and were significantly less in high cultch density treatment plots. • Field observations were more successful in detecting a recruitment event than planned recruitment detection devices. • Other environmental factors such as biofouling, algae coverage, predation, and competition with other oyster species may impact the ability of O. conchaphila to become a self‐sustaining population.

Chapter 3: Restoration in Practice Ecological restoration is defined as “The process of assisting the recovery of an ecosystem that has been degraded, damaged, or destroyed” (SERI 2004). This process includes multiple methods and techniques from ecological, restorationist, socioeconomic, cultural, educational, and political disciplines (Higgs 1997, Halle and Fattorini 2004). Expertise from these fields is useful in the development, implementation, and evaluation of restoration projects. Throughout this attempt to re‐establish Olympia oysters, several themes arose which encompass this broad range of disciplines. These themes parallel some of the restoration project guidelines presented by Clewell et. al. (2007). This chapter provides a synthesis of the Netarts Bay Olympia oyster restoration project, an overview of the lessons learned, and recommendations for this and other future restoration projects.

Defining long term restoration goals & a reference system Restoration ecologists recommend the establishment of restoration goals and a reference system as part of the initial planning process (Vivian‐Smith 2001, Clewell and Aronson 2007). Identifying a guiding image (sensu Palmer et al. 2005) will help with the development of criteria for implementation and effective monitoring. Reference systems have ecological characteristics similar to those desired at the restoration site (Brinson and Rheinhardt 1996). Some restoration projects may not have access to a reference site or may be faced with external factors which make the use of a reference site no longer feasible. When the use of a reference system is not possible, an alternative option is to develop restoration parameters and goals which guide researchers and practitioners through the restoration, providing benchmarks for comparison and assessment. In this instance, it remains extremely important to identify a long‐ term restoration goal and define what achieving “success” entails (Hobbs and Harris 2001, Zedler 2007). With clear project objectives and benchmarks, resources such as time and money can be used most effectively and science can be incorporated into the restoration design (see page 45).

The overall goal of the Netarts Bay oyster restoration is to re‐establish a self‐sustaining population of native oysters, which is one attribute of a restored system (SERI 2004). Specific goals regarding ideal densities of oysters and reef aggregate formations have not been established, in part due to the lack of knowledge of west coast oyster reefs (see page 5). We

43 were unable to identify a suitable reference site within the Bay or throughout Oregon. Instead, qualitative, historic descriptions from government publications, literature, and from books (Townsend 1893, Edmondson 1923, Dimick et al. 1941, Fitch 1953, Steele 1957, Marriage 1958, Hertlein 1959, Barrett 1963, Espy 1977, Kreag 1979) were utilized. To guide the restoration image, qualitative descriptions of current oyster populations (Baker et al. 1999, Harris 2004, Groth and Rumrill 2008, Heiman et al. 2008) also provided information on the oyster. Current population distributions of Olympia oysters, which describe habitat areas that they did not evolve with such as rocky rip‐rap (Heiman et al. 2008), were not used in developing the image. Knowledge from local oystermen who had experience with working with C. gigas and had seen O. conchaphila within their beds was extremely informative. This information, as well as general biological and ecological background (Chapter 1), helped in the development of the restoration strategy to recreate an intertidal reef aggregate environment which would eventually become a self‐sustaining population of native oysters. However, this experimental restoration is one such attempt at describing a species which has a broad intertidal range and habitat forms. Multiple outplanting strategies and techniques could be successful in re‐establishing Olympia oysters as the general structure and function of Olympia oyster reef assemblages is unknown. Other projects should aim to test other reef aggregate re‐establishment techniques to understand how the size, shape, and extent of reef structures impact Olympia oyster density, growth, and reproduction. For instance, one restoration goal may be to re‐establish oyster areas similar to those found in Elkhorn Slough, CA (see Heiman et al. 2008).

Identifying limiting factors This re‐establishment project was designed to address two potentially limiting factors for populations of O. conchaphila in Netarts Bay: inadequate broodstock and limited settlement substrate (Baker et al. 1999, Harris 2004, Brumbaugh et al. 2006). However, there may be other abiotic or biotic factors which may influence the spatial and temporal re‐establishment of the oysters (Temperton and Hobbs 2004). These factors may arise after project implementation, which is an important reason for continued monitoring and evaluation. As a restored population, the oysters will ideally be resilient (sensu Holling 1973) and able to overcome these limiting factors (SERI 2004, Palmer et al. 2005). Several potential limiting factors which could impact the effectiveness of the restoration (described below) were identified within the literature and at the re‐establishment site. Once limiting factors have been recognized, these

44 factors should be managed and accounted for before restoration continues, as a normal, healthy oyster population should be able to cope with these effects.

Water quality Current water quality threats include the presence of raw sewage, bacterial contamination, or marine fuel, all of which may be detrimental to O. conchaphila survival (Baker 1995). Additional water quality concerns include: low dissolved oxygen, chlorine treated sewage, non‐point source pollution, increasing eutrophication, sedimentation, siltation, herbicides, and trace metals (Cook et al. 1998, Shaffer 2004). Prolonged exposure to poor water quality conditions may cause mortalities, illnesses, or deformities in the filter feeders. It is important for oyster restoration projects to address current and potential water quality conditions to ensure the health and longevity of the oysters.

Predators, parasites, and pathogens Natural predators of the Olympia oyster include crabs, snails, sea stars, stingrays, leopard sharks, and aquatic ducks (Baker 1995). Nonnative predators also pose threats to O. conchaphila populations. The European green crab, Carcinus maenas, was initially documented in Netarts Bay in 2003. This may pose a threat to O. conchaphila because in laboratory experiments, the predation rates of C. maenas were highest on O. conchaphila compared to other bivalves (Palacios and Ferraro 2003). With the addition of C. maenas to the native predators, O. conchaphila may face increased predation pressure, and smaller oysters and recruits are more susceptible to predation due to their smaller size. Oyster drills prey on bivalves by drilling a hole into the shell and eating the organism from the inside out. There are two native drills, the thaids Nucella lamellosa and Acanthina spirata. O. conchaphila benefits from the presence of N. lamellose, which preys on other organisms (barnacles and mussels) and frees up habitat space otherwise occupied by them (Baker 1995). Nonnative drills have the potential to decimate oyster populations and were introduced when populations of nonnative oysters were imported for aquaculture purposes. These include Ocinebrellus inornatus (Buhle et al. 2005), Urosalpinx cinera (Atlantic oyster drill), and Cerastoma inornatum (formerly Ocenebra japonica, the Japanese oyster drill) (Baker 1995). C. inornatum (formerly O. japonica) caused high mortalities among the C. gigas commercial populations in Netarts Bay before the mid 1970’s (Kreag 1979). The flatworm Pseudosylochus ostreophagus (Japan) drills holes into juveniles, causing mortality before the oysters are able to reproduce. Although not directly

45 harmful to the oyster, Bryozoans which foul the shell substrate and oyster shells may compete with larvae for settlement space.

Parasites and pathogens are another cause for mortality and disease in O. conchaphila. Within San Francisco Bay, a recent study found several diseases and disease agents which may hinder recovery and restoration of the oyster (Friedman et al. 2005). Mikrocytos were discovered in O. conchaphila from Yaquina Bay and caused mortalities in O. edulis and C. gigas; no mortality studies were performed on O. conchaphila to determine potential effects of survival (Baker 1995). More complete details on parasites and pathogens are found in Baker (1995). Recent outbreaks of Vibrio spp. in Netarts Bay may inhibit reproduction and larval survival and has the capacity to kill larval C. gigas and O. edulis in the water column (Estes et al. 2004).

Aquaculture Netarts Bay is home to several commercial aquaculture beds where C. gigas is grown using bottom culture techniques. Restoration projects near the commercial beds may facilitate the spread of predators like the oyster drill, which would be detrimental to both oyster species. The re‐establishment cultch includes C. gigas (Figure 4, page Error! Bookmark not defined.; page 39), which can compete for space with O. conchaphila. Other project complications may arise if O. conchaphila spat settle on the C. gigas for commercial sale, potentially fouling the oysters and creating more work for the oystermen who have to remove the natives. Additionally, continual oyster harvest removes available substrate for potential spat settlement (Breitburg et al. 2000). This is of concern in the PNW because O. conchaphila settles on C. gigas and because C. gigas is continually removed, C. gigas is not a permanent reef structure in these settings. Permanent structures are more desirable such that multiple age classes and intricate physical structures could exist.

Incorporating science into the design and monitoring of the project Restoration ecologists have issued multiple pleas for projects to appropriately include science into the design and monitoring of restoration projects (e.g. Zedler 2000). Science‐driven restorations are based upon ecological principles, have distinct hypotheses and tests, therefore having rigorous results which can be applied to other systems. However, it is not always possible or practical for restoration projects to take a rigorous approach if large scale restoration

46 progress is to be made (Cabin 2007). Restoration projects may be required to use local and traditional knowledge with no scientific basis (Higgs 2005). A balance between these approaches is possible, where both restoration ecologists and restoration practitioners can apply a uniform, scientific process resulting in improved restoration knowledge (Giardina et al. 2007). Each restoration project is unique and requires its own assessment; however, there are procedures for restorationists which, when followed, can make a restoration more successful (Clewell et al. 2007). This project was also guided by a publication from The Nature Conservancy (TNC) and the National Oceanic and Atmospheric Administration (NOAA) Restoration Center, A Practitioners' Guide to the Design and Monitoring of Shellfish Restoration Projects (Brumbaugh et al. 2006).

This project was complicated by a permitting procedure which delayed outplanting and prevented the collection of pre‐implementation site data. However, the project was successfully implemented and captured important information related to oyster restoration. Setting up a randomized block experiment, although desirable for statistical purposes, was not feasible given the short amount of time and limited resources at hand. This project was fortunate to have a first year of intensive monitoring, but long term monitoring will be necessary to assess how well the oysters survived, how the eelgrass were impacted, and the examination of any other interactions within the bay. As monitoring resources are limited, there can either be frequent monitoring on a short‐term basis, or infrequent monitoring on a long‐term basis. Adaptive restoration, which utilizes monitoring data to inform management and restoration decisions, would benefit the most from long‐term data collection (Zedler and Callaway 2003). Guidelines for this level of restoration planning have been detailed by Short for seagrass and marsh systems and can be used as a template for this project (2000), along with additional oyster restoration criteria (Coen and Luckenbach 2000).

From this pilot study of the re‐introduction of Olympia oysters on a broad scale within Netarts Bay, we were able to come to several conclusions to aid in the development of future restoration plans. First, our results should be very encouraging to future research projects. These projects should expand upon these results to examine the relative survival, growth, and reproduction among multiple brood years across multiple cultch density treatments. Determining the minimum viable population necessary for the oysters to sustain themselves will also aid in the restoration efforts. Additional studies should examine how and when managers

47 can stop supplementing the oyster population with shell substrate (Mann 2007). Oyster restoration is more challenging because the shell substrate suffers shell degradation over time (or in the case of the eastern oyster, shell removal via harvest) (Powell et al. 2006). If the oyster birth rate does not exceed the death rate, there will be an insufficient number of cultch to allow for the population to expand (Powell and Klink 2007). Criteria and goals should be established for when the re‐establishment efforts stop the continued supplementation of shell substrate.

Policy: Long term management of Olympia oysters, eelgrass, and EFH The long term management of the resources will determine the ultimate success of the re‐establishment project. Current policies regarding harvest of Olympia oysters are prohibitive in Oregon, but interestingly, when told about the re‐establishment of the native oyster, most people ask when oysters will be available for consumption. Policies will eventually need to be developed to address this issue, especially for Netarts Bay as one of the most common recreational clamming destinations in Oregon. Other long‐term management considerations might involve the commercial oyster industry, particularly the aquaculture of C. gigas. Qualitative experiments on whether C. gigas and O. conchaphila compete for food sources would be useful in determining if the species are in direct competition with each other.

Long term management of eelgrass in the Pacific Northwest also will factor into the expansion of Olympia oyster restoration projects. Globally, seagrasses are experiencing declines (Orth et al. 2006). The goal of this study was to examine the impacts to eelgrass not only to investigate the system‐wise impacts of the restoration project, but also because there was a management issue with the potential adverse impact to eelgrass beds and subsequent EFH (see Chapter 1). Permitting of future Olympia oyster re‐establishment projects is subject to whether the eelgrass may be harmed, if the restoration project overlaps with EFH. Netarts Bay in particular contains the most robust eelgrass beds in Oregon, and as a “Conservation Estuary”, is considerably less impacted than other bays. Therefore, our results may not easily translate for other systems which are more degraded or impacted. Additionally, these results are from only one season but eelgrass can have relatively robust belowground biomass reserves, so the observed restoration impacts may only occur for a short term and the eelgrass abundances could rebound; or the eelgrass resources may slowly be exhausted as the bed attempts to rebound; or some other unforeseen result. We are unsure of what would happen if this re‐

48 establishment technique were to be applied in a system that is more impacted and how the impacts observed translate to other systems.

Another management consideration may be the incorporation of native oysters into the definition of essential fish habitat (EFH). Although the amount research on the Olympia oyster has been on the rise, there is still much left to be understood regarding the role of oysters as EFH. Research on the eastern oyster will aid in the understanding and management of the species. For instance, reefs of the native eastern oyster C. virginica are considered EFH (Coen et al. 1999), and although qualitative research has yet to be initiated for O. conchaphila, west coast oyster researchers consider Olympia oysters to be a part of EFH (Peterson et al. 2003, Ruesink et al. 2005).

Other important long term management considerations include the balancing of continued coastal development and other conservation pressures on the ecosystems (Thom et al. 2005). In addition to these changes, global climate change may impact the restoration of Olympia oysters. Even if populations of O. conchaphila are restored, they will still face unknown threats to their overall stability. Climate change poses the threat of more random and more severe disturbance events, rising temperatures, along with rising sea levels (International Panel on Climate Change 2001). It is possible that by 2100, restored oyster communities may no longer exist due to decreases in estuarine salinity (Scavia et al. 2002). Climate change may increase the degree of estuarine eutrophication, thereby increasing the amount of phytoplankton, which may be beneficial for oysters if they are capable of higher filtration rates. Alternatively, the higher algal loads in the water column may lead to anaerobic conditions as the algae decays. Changing water temperatures also have a potential to impact the distribution of the oyster and its reproductive timing, which is related to water temperature. These possible scenarios emphasize the importance of restoring resiliency in ecosystems.

Public involvement and restoration inertia For any restoration project to truly take hold and gain inertia, public interest and involvement in the project is key. Education and outreach activities are essential for building grassroots support of restoration activities, particularly with a project such as this which generated much excitement from academia and the general public. Once the general public becomes acquainted with the Olympia oyster, most people are very excited, positive, and

49 enthusiastic about its restoration (pers. obs.). Therefore, the biggest, most important challenge in restoring the Olympia oyster is reacquainting the Pacific Northwest with their native “Oly” and the stories and history which came along with the oyster. The PNW already has a strong attachment to salmon as a charismatic species, and the potential exists for another attachment to the Olympia oyster. There are several books and publications about our oyster heritage which can be used to increase outreach and education efforts (Gordon et al. 2001, Peter‐Contesse and Peabody 2005).

With increased interest in the plight of the Olympia oyster, interest in its restoration efforts may begin to gain momentum. Ecological restoration has a strong value‐based component (Higgs 1997, Davis and Slobodkin 2004, Clewell and Aronson 2007). In regions where residents value habitat restoration, projects are more likely to have public and financial support. Organizations like the Puget Sound Restoration Fund have successfully recruited the public general public and land‐owners for oyster restoration volunteer opportunities or spaces. NOAA’s Community‐Based Restoration Program is designed to develop stewardship through restoration activities. In the case of the restoration project in Netarts Bay, Netarts and nearby Oceanside are growing communities of retirees with extra time on their hands. This represents an untapped pool of potential community involvement, volunteer time, and word of mouth promotion for the project. There is the potential for untapped community support which could be integrated into future re‐establishment and monitoring protocols.

This re‐establishment effort is one of many initial Olympia oyster restorations underway on the west coast, as mentioned in Chapter 1. The overall restoration process is distinct in that a West Coast native oyster restoration working group was established and has held several workshops already. All participants have a common goal of restoring and re‐establishing the Olympia oyster and these workshops provide an effective means for communication between scientists, restoration practitioners, and other interested parties. Creating the TNC/NOAA West Coast Native Oyster Restoration Working Group represented an excellent method of communication and information sharing. This enabled all stakeholders, even if not part of the working group, to have access to this information and potentially use it for future projects. The public perception of research activities needs to remain positive to encourage volunteer participation and even volunteers who donate their land and/or leases toward the restoration

50 efforts. Additionally, there was significant interest expressed by academia and the general public in restoration of the native oyster, and these people could be used as volunteers.

Conclusions This work represents a balance between science of restoration ecology and the practice of ecological restoration (Weinstein 2007). It was important to test multiple restoration methods while ensuring the project retains applicability to future restoration efforts. There is a constant challenge to balance restoration needs and scientific needs, all within the greater context of conservation (Cabin 2007). Netarts Bay will most likely always have a C. gigas presence, and restoration will need to account for this fishery presence (Weinstein 2007).

Restoration is complicated regardless of the site location. In this case, there was no reference site, which immediately made our project goals more complicated and difficult to define, as opposed to those projects which establish reference sites and strive for fidelity (Higgs 1997, p. 343). There is still have much to learn about the biology and ecology of the native oyster, which will aid in the restoration process. Proceeding without a decent understanding of the oyster may eventually lead to project complications. Restoration must take place on the ecosystem scale. Although the oysters are the focus of this project, it is important to take the greater estuarine habitat dynamics into account when attempting to interpret oyster growth and reproductive habits. There are many other factors to monitor in the bay which may impact the re‐establishment project, such as algae populations, predation by native and non‐native predators, and water quality for temperature and Vibrio spp. populations.

Although Olympia oyster restoration projects are in their infancy, it is important for management agencies and interest groups to begin long‐term planning for the ultimate recovery of the species. This includes defining “recovery” itself, addressing Essential Fish Habitat, harvest of the species, and greater ecosystem interactions. It is also important for the organizations conducting the restoration to increase education and outreach if they wish to increase community support and involvement.

Chapter 4: Concluding thoughts Olympia oyster restoration is gaining momentum along the North American West Coast. The objective of this thesis was to build on previous studies and to provide researchers, practitioners, and other interested parties with a synthesis of information which can be used to guide future projects. There is a lack of qualitative information about the ecology of Olympia oysters, which this study aimed to reduce. The availability of qualitative information supplements the detailed scientific information and enables restorationists to make informed decisions on where and how to restore the native oyster. Early 20th century laboratory studies provide a scientific basis for the biology of the oyster, while cultural accounts of early PNW oystering aid in the creation of an image of what the oysters could be restored to. Government surveys and fishery reports lend clues as to the abundance and harvest of the oysters, where lower harvests indicated a decline in the abundance of the oyster. Olympia oyster restoration and re‐establishment projects can also look to the East Coast native oyster, C. virginica, for guidance and knowledge of reef structure restoration. Although the oysters have differing biology and ecology, the work done on C. virginica still and principles developed for restoration are still applicable to the Olympia oyster.

The contributions of this study to Olympia oyster restoration are detailed information of monthly growth for year one and year two oysters. This study demonstrated Olympia oysters are capable of surviving and reproducing in an area of their former range, but more detailed information is needed on the reproductive and recruitment cycles. The cultch density treatments used in this re‐establishment attempt may or may not represent the “reef aggregates” characteristic of Olympia oysters; recommendations to further test and develop reef structures were included with this document. The response of eelgrass to cultch treatments was of particular interest, as results indicated eelgrass abundance declined in treatment plots with high cultch densities (19%). There were fundamental differences between the eelgrass bed locations, although they were in close proximity to one another. Additional ecological interactions were found to influence eelgrass beds at the site, such as macroalgae abundance, but the cultch treatment effect was observed before the macroalgae boom. A consistent pattern was observed where eelgrass declined as cultch density increased, even after variation due to location and month were accounted for. The low and medium density cultch plots (4% and 11% respectively) did not have statistically significant eelgrass declines; further testing of oyster and

52 eelgrass abundances at these densities will provide greater insight as to the best possible habitat for oyster re‐establishment.

Continued monitoring and future research are always included as recommendations for restoration projects and this particular effort makes no exception. Restoration of a native species is not an immediate process and only with patience, attention to detail, and continued efforts will the full restoration of Olympia oysters be realized. Long‐term monitoring of oysters and eelgrass at the Netarts Bay site will aid in the understanding of oyster aggregate reef recovery. Experimental‐based restorations will allow researchers to develop sound conclusions and provide other restoration groups with information to utilize and build upon. It is important for Olympia oyster restoration work to be applicable to other estuaries and environments, as these contributions can increase the efficacy of the re‐establishment of the species as a whole. Research on Olympia oyster re‐establishment may also be used to shape policies governing the management of native oysters and the eelgrass beds they inhabit. In particular, the definition of EFH may be impacted by the relationship between native oysters and the native eelgrass of the PNW. Above all, it will be important to continue education and outreach not only to other research groups and government, but to the general public who live in the PNW. Increasing awareness of the native oyster upon which the PNW economy was founded is likely to increase the demand for restoration projects and perhaps even public incentives to become involved with the re‐establishment of the beloved “Oly.”

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Appendices

Appendix A. ANOVA tables for 2005 oyster shell length. Type III Sum of Source Squares df Mean Square F Sig. Location 53.335 1 53.335 0.231 0.632 Month 68438.536 5 13687.707 59.353 <0.001 Shelltrt 1065.579 2 532.789 2.310 0.107 Location * Month 274.620 5 54.924 0.238 0.944 Location * Shelltrt 475.157 2 237.578 1.030 0.363 Month * Shelltrt 1597.317 10 159.732 0.693 0.728 Month * Location * Shelltrt 1287.878 10 128.788 0.558 0.842 Error 15451.167 67 230.614 Total 369564.000 103

Appendix B. Table of oyster reproductive activity by month. Month May Jun(1) Jun(2) Jul Aug Active 19 26 17 15 22 Larvae 12100 Non-active 32282 Total 23 30 20 23 24

Appendix C. ANOVA table for oyster density. Type III Sum of Source Squares df Mean Square F Sig. Location 0.017 1 0.017 0.188 0.666 Month 0.116 4 0.029 0.316 0.866 Cultchtrt 4.353 2 2.176 23.660 <0.001 Location * Month 0.754 4 0.188 2.049 0.100 Location * Cultchtrt 0.008 2 0.004 0.044 0.957 Month * Cultchtrt 0.794 8 0.099 1.079 0.391 Location * Month *Cultchtrt 1.233 8 0.154 1.676 0.125 Total 24.148 86

Appendix D. Mean daily temperatures at experimental site. Mean daily temperatures (°C) for re‐establishment site. Mean of six loggers.

Appendix E. Salinity measured at site. Salinity as measured by CTD during site visits on both outgoing and incoming tides.

28 Salinity

20 4/28 5/18 6/7 6/27 7/17 8/6 8/26 9/15 10/5 Date, 2007

Appendix F. ANOVA details for eelgrass parameters. Eelgrass parameter ANOVA interaction terms and Games‐Howell Multiple Comparisons for the factor of month.

Eelgrass parameter Significant Interactions Games‐Howell Multiple Comparisons

Percent cover None April ≠ June, April ≠ July

Shoot density Location * Month May ≠ July, June ≠ July Location * Month, Flowering shoot % Cultch Treatment * Location May ≠ June, June ≠ July April ≠ May, April ≠ June, April ≠ July, Blade length None May ≠ June, May ≠ July

Blade width None None

Appendix G. Mean macroalage percent cover. Macroalgae percent cover as by cultch treatment and location, shown with standard deviations.

Cultch Treatment and Location Control Low Medium High Month ABABABAB April00000000 Standard Dev.00000000 May00000000 Standard Dev.00000000 June 2.9 0 6.1 0 26.7 10 24.4 27.2 Standard Dev. 3.44 0 4.19 0 28.48 15.9 16.69 42.82 July0000.6000.61.1 Standard Dev.0 0 00.960 00.961.92 August 0 0 ‐‐‐‐00 Standard Dev. 0 0 ‐‐‐‐00 September ‐‐003.3000 Standard Dev. ‐‐0 0 5.77 0 0 0

Appendix H. Table of means for eelgrass parameters of percent cover and shoots m‐2. Percent cover Total shoots Treatment Location Mean Stdev SE Mean Stdev SE Control A 73.8 16.73 4.83 692.9 298.30 86.11 Low A 89.0 7.09 2.05 720.1 225.42 65.07 Medium A 69.8 14.52 4.38 561.2 174.34 52.57 High A 58.5 12.77 3.85 441.8 140.85 42.47 Control B 88.6 5.72 1.65 463.7 74.08 21.38 Low B 84.7 9.84 2.84 391.6 85.91 24.80 Medium B 75.8 17.83 5.38 357.1 108.62 32.75 High B 66.8 26.23 7.91 340.7 99.91 30.12

Appendix I. Graphs of eelgrass blade length. By location ± SE.

100 (a) Location A 100 (b) Location B

80 80 (cm)

60 60 Length

Blade 40 40

20 20 Eelgrass

0 0 April May June July April May June July

Control Low Medium High Control Low Medium High

Appendix J. ANOVA table for eelgrass blade length. Type III Sum of Source Squares df Mean Square F Sig. Location 0.482 1 0.482 44.422 <0.001 Month 1.231 3 0.410 37.784 <0.001 Cultchtrt 0.218 3 0.073 6.688 0.001 Location * Month 0.018 3 0.006 0.566 0.639 Location * Cultchtrt 0.061 3 0.020 1.865 0.145 Month * Cultchtrt 0.023 9 0.003 0.231 0.989 Location * Cultchtrt * 0.045 9 0.005 0.462 0.894 Month Error 0.652 60 0.011 Total 251.007 92

Appendix K. Graphs of eelgrass blade width. Monthly means ± SE.

10 10

(mm) 8 8

6 6 width

4 4 blade

2 2

Eelgrass (a) Location A (b) Location B 0 0 May June July May June July

Control Low Medium High Control Low Medium High

Appendix L. ANOVA table for eelgrass blade width. Type III Sum of Source Squares df Mean Square F Sig. Location 0.151 1 0.151 33.459 <0.001 Month 0.009 2 0.004 0.946 0.397 Cultchtrt 0.057 3 0.019 4.167 0.011 Location * Month 0.003 2 0.001 0.314 0.733 Location * Cultchtrt 0.021 3 0.007 1.524 0.222 Month * Cultchtrt 0.013 6 0.002 0.487 0.814 Location * Cultchtrt * Month 0.018 6 0.003 0.677 0.669 Error 0.190 42 0.005 Total 38.802 66