Parasites and Disease in Oysters and Mussels of U.S. Coastal Waters

National Status and Trends, the Mussel Watch Monitoring Program

NOAA National Centers for Coastal Ocean Science Center for Coastal Monitoring and Assessment D. A. Apeti Y. Kim G.G. Lauenstein E.N. Powell J. Tull R. Warner

December 2014 NOAA TECHNICAL MEMORANDUM NOS NCCOS 182 NOAA NCCOS Center for Coastal Monitoring and Assessment CITATION

Apeti, D.A., Y. Kim, G.G. Lauenstein, E.N. Powell, J. Tull, and R. Warner. 2014. Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters. National Status and Trends, the Mussel Watch Monitoring Program. NOAA Tech­ nical Memorandum NOSS/NCCOS 182. Silver Spring, MD 51 pp.

ACKNOWLEDGEMENTS

The authors would like to acknowledge Juan Ramirez of TDI-Brooks International Inc., and David Bushek and Emily Scarpa of Rutgers University Haskin Shellfish Laboratory for a decade of analytical effort in providing the Mussel Watch histopathology data. We also wish to thank Erik Davenport, who provided data analysis support, and most importatntly reviewers Gretchen Messick and Shawn McLaughlin of the NCCOS Cooperative Oxford Laboratory, as well as Kevin McMahon and Greg Piniak, for their invaluable assistance in making this document a better product than what we had initially envisioned.

Mention of trade names or commercial products does not constitute endorsement or recommendation for their use by the United States government. Parasites and Disease in Oysters and Mussels of U.S. Coastal Waters.

National Status and Trends, the Mussel Watch Monitoring Program

National Oceanic and Atmospheric Adminstration (NOAA) National Ocean Service (NOS) National Centers for Coastal Ocean Science (NCCOS) Center for Coastal Monitoring and Assessment (CCMA)

Authors Dennis A. Apeti, Gunnar G. Lauenstein, and Rob Warner NOAA Center for Coastal Monitoring and Assessment Coastal and Oceanographic Assessment, Status and Trends Branch

Yungkul Kim School of Science, Engineering & Mathematics Bethune-Cookman University

Eric N. Powell Gulf Coast Research Laboratory University of Southern Mississippi

Jamila Tull School of the Environment Florida A&M University

NOAA Technical Memorandum NOS NCCOS 182 December 2014

United States Department National Oceanic and Atmospheric National Ocean Service of Commerce Administration

Penny Pritzker Kathryn Sullivan Holly Bamford Secretary Under Secretary Assistant Administrator

NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program TABLE OF CONTENTS

Acronyms ...... ii List of Tables ...... iii List of Figures ...... iii Executive Summary ...... vii Introduction ...... 1 Table of Contents Table Methods ...... 3 Monitoring Sites and Bivalve Species ...... 3 Sample Collection ...... 4 Analytical Methods ...... 4 Data Analysis and Statistical Approach ...... 7 Results ...... 10 Parasites...... 10 Prokaryotic Inclusion Bodies ...... 10 Gregarines ...... 12 Haplosporidium nelsoni ...... 14 Perkinsus marinus ...... 16 Ciliates ...... 18 Cestodes ...... 20 Trematodes ...... 22 Nematodes ...... 24 Copepods...... 26 Pinnotherid Crabs ...... 28 Disease...... 30 Tissue Hemocytic Infiltration ...... 30 Ceroid Bodies ...... 32 Digestive Gland Atrophy ...... 34 Tissue Necrosis ...... 36 Xenomas ...... 38 Synthesis ...... 40 Conclusion ...... 46 Literature Cited...... 47 Appendix A...... 51 Appendix B...... 52

i NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program ACRONYMS AND TECHNICAL TERMS

Ag – silver American oyster– Crassostrea virginica also known as Eastern oyster As – arsenic Blue mussel – Mytilus edulis (East Coast), M. californianus, M. galloprovincialis (California coast), M. trossulus (Northern California to Alaska) CCMA – Center for Coastal Monitoring and Assessment Cd – cadmium Cestode – a parasitic flatworm with a specialized organ for attachment

Table of Contents Table COAST – Coastal Ocean Assessments, Status, and Trends Branch Cr – chromium Cu – copper DDT – dichlorodiphenyltrichloroethane Dermo – a common name for an oyster disease caused by Perkinsus marinus, a parasitic pathogen that affects oysters Hawaiian oyster – Ostrea sandvicensis Hg – mercury Intensity - a measure of severity of infection or disease in infected or diseased organisms MHE stain - Meyer’s hematoxylin and eosine stain Mn – manganese MSX – multinucleated sphere X/unknown, an oyster disease caused by Haplosporidium nelsoni MWP – Mussel Watch program NCCOS – National Centers for Coastal Ocean Science Necrosis – tissue death Nematode – a roundworm with an unsegmented body Ni – nickel NOAA – National Oceanic and Atmospheric Administration NOS – National Ocean Service NS&T – National Status and Trends PAH – polycyclic aromatic hydrocarbons Pb – lead PCB – polychlorinated biphenyls PIB – prokaryotic inclusion bodies Prevalence - the proportion of individuals in a sample population that are infected or diseased at a given time Se – selenium Sn – tin Xenoma – an enlarged lesion composed of hypertrophic host cells filled with ceratin parasites, such as ciliates; a cyst Zebra mussel – two species of invasive mussels found in the Great Lakes, Dreissena polymorpha (zebra mussel) and D. bugensis (quagga mussel) Zn – zinc

ii NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program LIST OF TABLES

Table 1. List of organic pollutants and metals analyzed by the NS&T Program. Table 2a. List of parasites measured by MWP in sentinel bivalves. Table 2b. List of disease and host responses measured by the MWP in sentinel bivalves. Table 3a. Semi-quantitative scale for Haplosporidium nelsoni (MSX) infection modified from Ford and Figueras (1988). Table 3b. Semi-quantitative scale for Perkinisus marinus [adapted from Mackin (1962) by Craig et al. (1989)]. Table 3c. Semi-quantitative scale for trematode sporocyst. Table of Contents Table Table 3d. Semi-quantitative scale for digestive gland atrophy. Table 4. Average prevalence (%) of parasites and diseases by region and by bivalve type (nd indicates that parameter was not detected in a given bivalve species) based on the 2008 - 2009 MWP data. Table 5. Average intensity of parasite infections and diseases by region and by bivalve type (nd indicates that parameter was not detected in a given bivalve species) based on the 2008 - 2009 MWP data. Table 6. Comparison of prevalence of histopathology parameters among bivalve species from 2008 - 2009 using analysis of variance (ANOVA). For each parameter, bivalves that do not have the same letter are statistically differ­ ent at p < 0.05 (nd = parameter was not detected). Table 7. Comparison of prevalence of histopathology parameters in bivalve among coastal regions from 2008 - 2009 using analysis of variance (ANOVA). For each parameter, coastal regions that do not have the same letter are statisti­ cally different at p < 0.05 (nd = parameter was not detected).

LIST OF FIGURES

Figure 1. National distribution of Mussel Watch program sampling sites, coded to reflect the species collected there. Figure 2. Prokaryotic inclusion bodies (PIB) in digestive tract epithelium of an American oyster. Arrows indicate PIBs (Kim et al., 2006). Figure 3. Spatial distribution of prokaryotic inclusion bodies (chlamydia and rickettsia) prevalence in oysters and mus­ sels, based on 2008 - 2009 Mussel Watch data. Figure 4. Spatial distribution of prokaryotic inclusion bodies (chlamydia and rickettsia) infection intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 5. Site-specific long-term temporal trends of prokaryotic inclusion bodies (chlamydia and rickettsia) prevalence in oysters and mussels. Figure 6. Section of oocyst (arrow) of gregarine Nematopsis sp. in the connective tissues of scallop (left); in oyster (right) (Meyers and Burton, 2009). Figure 7. Spatial distribution of gregarine prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 8. Spatial distribution of gregarine infection intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 9. Site-specific long-term temporal trends of gregarine prevalence in oysters and mussels. Figure 10. Numerous multinucleated plasmodia of Haplosporidium nelsoni in the gills of an American oyster. Arrows point to example parasites (Kim et al., 2006). Figure 11. Spatial distribution of H. nelsoni prevalence in oysters, based on 2008 - 2009 Mussel Watch data. Figure 12. Spatial distribution of H. nelsoni infection intensity in oysters, based on 2008 - 2009 Mussel Watch data. Figure 13. Site-specific long-term temporal trends of H. nelsoni prevalence in oysters.

iii NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Figure 14. Several Perkinsus marinus cells in the intestine of an American oyster (40x mag, MHE stained). Photo by Joe Marcino, Maryland Department of Natural Resources. Figure 15. Spatial distribution of P. marinus prevalence in oysters, based on 2008 - 2009 Mussel Watch data. Figure 16. Spatial distribution of P. marinus infection intensity in oysters, based on 2008 - 2009 Mussel Watch data. Figure 17. Site-specific long-term temporal trends of P. marinus prevalence in oysters. Figure 18. Ciliates in the intestine of an oyster (left) and between gill filaments of a blue mussel (right) (Kim et al., 2006). Figure 19. Spatial distribution of ciliate prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch data.

Table of Contents Table Figure 20. Spatial distribution of ciliate infection intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 21. Site-specific long-term temporal trends of ciliate prevalence in oysters and mussels. Figure 22. Encapsulated cestode larvae in the vesicular connective tissue surrounding the stomach of an American oyster (Kim et al., 2006). Figure 23. Spatial distribution of cestode prevalence in oysters, based on 2008 - 2009 Mussel Watch data. Figure 24. Spatial distribution of cestode infection intensity in oysters, based on 2008 - 2009 Mussel Watch data. Figure 25. Site-specific long-term temporal trends of cestode prevalence in oysters. Figure 26. Trematode sporocyst (Bucephalus sp.) infection in blue mussels (Meyers and Burton, 2009). Figure 27. Spatial distribution of trematode prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 28. Spatial distribution of trematode infection intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 29. Site-specific long-term temporal trends of trematode prevalence in oysters and mussels. Figure 30. Unidentified nematode larvae in connective tissue of the digestive gland of an American oyster (Kim et al., 2006). Figure 31. Spatial distribution of nematode prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 32. Spatial distribution of nematode infection intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 33. Site-specific long-term temporal trends of nematode prevalence in oysters and mussels. Figure 34. Copepod (*) in gill surrounded by an intense hemocyte reaction (Carballal et al., 2001). Figure 35. Spatial distribution of copepod prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 36. Spatial distribution of copepod infection intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 37. Site-specific long-term temporal trends of copepod prevalence in oysters and mussels. Figure 38. A pea crab (Pinnotheridae family) living inside an American oyster (Howard et al., 2004). Figure 39. Spatial distribution of pinnotherid crab prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 40. Spatial distribution of pinnotherid crab infection intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 41. Site-specific long-term temporal trends of pinnotherid crab prevalence in oysters and mussels. Figure 42. Hemocytic infiltration (arrow) near the gill base of an American oyster (Kim et al., 2006). iv NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Figure 43. Spatial distribution of hemocytic infiltration prevalence in oysters and mussels, based on 2008 - 2009 Mus­ sel Watch data. Figure 44. Spatial distribution of hemocytic infiltration intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 45. Site-specific long-term temporal trends of tissue hemocytic infiltration prevalence in oysters and mussels. Figure 46. Ceroid bodies in gonad tissue of an American oyster (40x mag, MHE stained). Photo by Joe Marcino, Maryland Department of Natural Resources. Figure 47. Spatial distribution of ceroid body prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 48. Spatial distribution of ceroid body intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. of Contents Table Figure 49. Site-specific long-term temporal trends of ceroid body prevalence in oysters and mussels. Figure 50. Atrophy (red arrow) of the digestive gland in an American oyster characterized by atrophic epithelia result­ ing in rounded and enlarged lumina compared to quadriradiate lumina (black arrow) formed by normal thick epitheli­ um (Kim et al., 2006). Figure 51. Spatial distribution of digestive gland atrophy prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 52. Spatial distribution of digestive gland atrophy intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 53. Site-specific long-term temporal trends of digestive gland atrophy prevalence in oysters and mussels. Figure 54. Necrosis in intestine of an American oyster (20x mag, MHE stain). Cells with pyknotic nuclei (arrow). Pho­ to by Joe Marcino, Maryland Department of Natural Resources. Figure 55. Spatial distribution of tissue necrosis prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 56. Spatial distribution of tissue necrosis intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 57. Site-specific long-term temporal trends of tissue necrosis prevalence in oysters and mussels. Figure 58. Xenomas filled with ciliates in gills of the mangrove oyster in earlier (left) and more advanced stage (right). Arrows = ciliates; arrowheads = nucleus of the host cell (Boehs et al., 2009). Figure 59. Spatial distribution of xenoma prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 60. Spatial distribution of xenoma intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data. Figure 61. Site-specific long-term temporal trends of xenoma prevalence in oysters and mussels. Figure 62. Parasite taxa richness in different host bivalve types by region. The upper and bottom limits of the boxplots represent the 25th and 75th percentiles, the whiskers represent the 5th and 95th percentiles, and the line in the middle of the box is the median. Figure 63a. Temporal variation of parasite taxa richness in oysters from the East Coast and the Gulf of Mexico from 1995 - 2009. Figure 63b. Temporal variation of parasite taxa richness in blue mussels from the East and West Coasts of the U.S from 1995 - 2009.

v NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Table of Contents Table

vi NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program EXECUTIVE SUMMARY

As a part of the National Centers for Coastal and Ocean Science, the National Status and Trends (NS&T) Mussel Watch program (MWP) uses bivalve mollusks (oysters and mussels) as sentinel organisms to monitor the health of our nation’s coastal and marine waters, including Alaska, Hawaii, Puerto Rico, and the Great Lakes. The program, which began in 1986 with 145 monitoring sites, today encompasses over 300 coastal locations nationwide, where bivalves are collected and measured for contaminant concentrations (organic compounds and metals). In 1995, histopathology was added to the program to survey the presence of parasites and disease in the mollusks. An array of parasites and disease (pathology) (e.g. cestodes, nematodes, trematodes, edema and necrosis) are quantified using direct quantitative counts or semi-quantitative scales.

This report, which is based on the 1995 - 2009 MWP monitoring data, provides both national and region­ Executive Summary al trends and distribution of parasites and disease in mussels and oysters. Because some of these sentinel mus­ sels and oysters hold important commercial and recreational value in the U.S., this first-ever national assess­ ment of MWP histopathology monitoring data provides coastal managers and the public with an overview of the most recent (2008 - 2009) conditions, and historical trends, of parasites and disease in these shellfish.

Results presented in this document indicate that prevalence (the measure of occurence), as well as intensity (the measure of severity), of parasites and disease varied broadly among host bivalves and geographic locations. In gen­ eral, prevalence and intensity ranged from low to medium. However, the presence of some hot-spots, with ele­ vated parasite infection and disease, were observed in some coastal areas. Oysters from the Gulf of Mexico had significantly higher parasite taxa than oysters from the East Coast, which in turn harbored more parasite taxa than oysters from Puerto Rico or Hawaii. Among the mussels, the West Coast mussels had significantly higher parasite taxa richness than East Coast mussels, which harbored more parasite taxa than mussels from Alaska. Zebra mussels, which are invasive in the Great Lakes, harbored few parasites and had significantly fewer cases of disease than all the other bivalves assayed. Many parasites and certain disease were only found in oysters and blue mussels. Tempo­ ral trends indicated that prevalence of parasites and disease were largely static and fairly minor over the monitoring period. Prevalence, intensity, and trends for both disease and parasites were low in the bivalves during this study.

Previous publications based on preliminary MWP data (1995 - 1998) noted, at regional levels, linkages be­ tween water temperature and occurrence of parasites (Kim and Powell, 2007), as well as positive correlations be­ tween pathology and contaminant concentrations (Kim et al., 2008). However, correlations between the his­ topathology parameters and contaminant body burdens in the bivalves were generally weak in the present study. The lack of correlation with high levels of contaminant body burden suggests that the cause may lie elsewhere.

For management purposes, this report constitutes an unprecedented baseline dataset against which his­ topathology measures could be evaluated during unforeseen events. As the only continuous coast­ al monitoring program that is national in scope, the MWP provides a unique and unparalleled opportu­ nity to determine national, regional and local perspectives of the health of our coastal ecosystems. This baseline information will be vital information for determining degrading conditions, as environmental stressors, including unforeseen natural and anthropogenic events and climate change, continue to impact coastal resources.

vii viii Parasites and Disease in Oysters and Mussels of U.S. Coastal Waters

INTRODUCTION

Coastal and estuarine environments contain diverse and burdens in bivalves have some linkage, the MWP began unique habitats that support biologically diverse estuarine surveying them for histopathology parameters in 1995, and marine species, which in turn support important rec­ with even earlier work performed on selected samples. reational and commercial fisheries. However, coastal and Parameters measured include an array of parasites, and Introduction marine waters are also a sink for organic and inorganic host responses to unknown stressess that cause pathology pollutants, which potentially have direct adverse effects (Tables 2a and 2b). on the habitats and biota, and indirect impacts on hu­ mans (through the food chain) who consume some of In this report, data from 1995 - 2009 were used to pro­ these marine resources. vide national and regional insight into the distribution and temproal trends of parasites and disease in mussels and The National Status and Trends Mussel Watch program oysters from US coastal waters, including the Great Lakes. (NS&T MWP) was designed to monitor the status and trends of chemical contamination in U.S. coastal waters and the health of sentinel shellfish (oysters and mussels) as indicators of water quality. The program was estab­ lished in response to a legislative mandate under Section 202 of Title II of the Marine Protection, Research and Sanctuaries Act (MPRSA) (33 USC 1442), which called on the Secretary of Commerce to, among other activities, initiate a continuous monitoring program. The MWP began in 1986 and is one of the longest running con­ tinuous coastal monitoring programs that is national in scope. The MWP is based on the collection and analy­ sis of sediments and bivalve mollusks. The MWP cur­ rently includes over 300 sites and measures more than 150 chemical pollutants, including: polycyclic aromatic hydrocarbons (PAHs); chlorinated pesticides, including dichlorodiphenyltrichloroethane (DDT) and its break- down products; tributyltin and its break-down products; polychlorinated biphenyls (PCBs); and trace elements (Table 1).

Mussels and oysters are sessile (stationary) filter-feeder organisms that are exposed to a variety of stressors, such as adverse physical conditions, and chemical and biologi­ cal toxins of the surrounding water. Thus, they are known to be good integrators of biological toxins and contam­ inants in a given area (Berner et al., 1976; Farrington et al., 1980; Farrington, 1983; Tripp and Farrington, 1984).

Bivalves can be exposed to parasites and disease as a di­ rect consequence of filter-feeding. Evidence of relation­ ships between certain tissue pathologies and contaminant exposure, as well as influence of contaminant exposure on bivalve defense mechanisms, has been established (Weis et al., 1995; Johnson et al., 1992; MacKenzie et al., 1995). Because pollutants, pathology, and parasite body

1 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Introduction

Oyster aquaculture (Image: NOAA National Marine Fisheries Service).

2 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

METHODS

Monitoring Sites and Sentinel Species

Mussel Watch sites were selected to represent large coastal areas that can be used to construct a nationwide Methods assessment. Mussel Watch sampled 145 sites nationwide in 1986. New sites were gradually added to the program, resulting in over 300 total sites today (Figure 1). Because a single species of mussel or oyster is not common to all coastal regions, several bivalve species were collected to gain a national perspective.

The American oyster, Crassostrea virginica (Gmelin, 1791), was sampled from coastal and estuarine areas of the East American oysters (Crassostrea virginica) Coast and the Gulf of Mexico. The mangrove oyster, Crassostrea rhizophorae (Guilding, 1828), was collected in Puerto Rico, and the Hawaiian oyster, Ostrea sandvicensis (Sowerby, 1871), was sampled in Hawaii. In this report, the inclusive general term “oyster” is used for all three.

Mytilid mussels, the blue mussel speices, were collect­ ed along the Northeast and West Coasts. Mytilus edulis (Linnaeus, 1758) were collected on the Northeast Coast. Along the West Coast, different mytilid species are found. M. californianus (Conrad, 1837) were collected in southern California. However, from northern Califor­ nia to the coast of Washington, M. californianus, M. gallo- provincialis (Lamarck, 1819), and hybrid species between M. galloprovincialis and M. trossulus (Hilbish et al., 2000) are found and form a mytilids specis complex. In Alas­ ka, M. trossulus (Hilbish et al., 2000) were collected. For Blue mussels (Mytilus edulis) the purpose of this report, the West Coast species com­ plex (Mytilus spp.), as well as the East Coast Mytilus edulis, were designated by the common name “blue mussel.”

Zebra mussels, Dreissena polymorpha (Pallas, 1771), and quagga mussels, Dreissena bugensis (Andrusov, 1897), were sampled in the Great Lakes. D. bugensis and D. polymorpha were introduced from Europe, likely in ship ballast wa­ ter (Hebert et al., 1989). The MWP does not distinguish between the two speceis, however, detailed taxanomy of the two species are discussed by Rosenberg and Ludy­ anskiy (1994). In this report, the name “zebra mussels” is used in reference to both D. bugensis and D. polymorpha.

Zebra mussels (Dreissena polymorpha)

3

NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Methods

Figure 1. National distribution of Mussel Watch program sampling sites, coded to reflect the species collected there.

Sample Collection Analytical Methods

The standard operational procedures for Mussel Watch Contaminant Analysis bivalve collection were described in detail in Apeti et al. (2011) and Lauenstein and Cantillo (1998). Except in The MWP analyzes a broad suite of chemical con­ the Great Lakes, bivalve sampling occured during winter taminants that bioaccumulate in bivalve tissues, includ­ months, since reproduction related changes in lipid lev­ ing: PAHs; chlorinated pesticides, including DDT, and els during warmer months may affect contaminant body its break-down products; tributyltin and its break-down burden measurements (Jovanovich and Marion, 1987). In products; PCBs; and trace elements (Table 1). As a part general, bivalves were dredged, tonged, or handpicked in of the program’s quality control protocols, chemical anal­ intertidal or subtidal zones. In the Great Lakes, zebra mus­ yses follow stringent procedures, detailed in Kimbrough sels were collected by free-diving around mid-September, and Lauenstein (2006) for metals, and Kimbrough et al. since the lakes freeze in the winter. Sampling frequency (2007) for organic compounds. Lauenstein and Cantillo has varied over the years, but remained at a biennial fre­ (1998) also provide a detailed account of the analytical quency during the period of this study (1995 - 2009). For methods quality assurance protocols. The NS&T Pro­ histopathology analyses, about 20 oysters and blue mus­ gram reports contaminant concentrations on a dry weight sels, and 60 zebra mussels (due to their small size) were basis and the data are available at: http://egisws02.nos. collected at each monitoring site. After collection, sam­ noaa.gov/nsandt/#.Histopathology Analysis ples were preserved on ice and shipped within 48 hours to the analytical laboratory. All histopathology analyses were conducted at the Rut­ gers University’s Haskin Shellfish Laboratory. The labo­

4 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Table 1. List of organic pollutants and metals analyzed by

Metals: Silver (Ag), Arsenic (As), Cadmium (Cd), Chromium (Cr), Copper (Cu), Iron (Fe), Lead (Pb), Mercury (Hg), Manganese (Mn), Nickel (Ni), Selenium (Se), Tin (Sn), Zinc (Zn)

Butyltins: monobutyltin, dibutyltin, tributyltin, tetrabutyltin Methods Chlordanes: alpha-chlordane, gamma-chlordane, oxychlordane, cis-nonachlor, trans-nonachlor, heptachlor, Hep- tachlor-Epoxide Chlorpyrifos DDTs: ortho and para forms of parent 2,4’DDT and 4,4’DDT and metabolites 2,4’DDE; 4,4’DDE; 2,4’DDD; 4,4’DDD Dieldrins: aldrin, dieldrin and endrin Chlorobenzenes: 1,2,3,4-Tetrachlorobenzene, 1,2,4,5-Tetrachlorobenzene, Hexachlorobenzene, Pentachloroben­ zene, Pentachloroanisole Hexachlorocyclohexanes (HCHs): alpha-Hexachlorocyclohexane, beta-Hexachlorocyclohexane, delta-Hexachloro­ cyclohexane, gamma-Hexachlorocyclohexane Endosulfans: Endosulfan I, Endosulfan II, Endosulfan sulfate PAHs: Biphenyl, Naphthalene, C1-Naphthalenes, C2-Naphthalenes, C3-Naphthalenes, C4-Naphthalenes, Acenaphthylene, Acenaphthene, Fluorene, C1-Fluorenes, C2-Fluorenes, C3- Fluorenes, Anthracene, Phenan­ threne, C1-Phenanthrenes/Anthracenes, C2-Phenanthrenes/Anthracenes, C3-Phenanathres/Anthracenes, C4-Phenanthrenes/Anthracenes, Pyrene, Dibenzothiophene, C1-Dibenzothiophenes, C2-Dibenzothiophenes, C3-Dibenzothiophenes, Fluoranthene, C1- Fluoranthenes/Pyrenes, Benz[a]anthracene, Chrysene/Triphenyl­ ene, C1-Chrysenes, C2-Chrysenes, C3-Chrysenes, C4-Chrysenes, Benzo[b]fluoranthene, Benzo[jk]fluoran­ thene, Benzo[e]pyrene, Benzo[a]pyrene, Dibenzo[ah]anthracene, Benzo[ghi]perylene and Indeno[123-cd] pyrene ratory uses stringent quality control through blind assays The cross-section was immediately transferred to a tissue to correct bias and assure data quality. Detailed analytical capsule and placed in Davidson’s fixative for two days, fol­ protocols are provided in the NOAA Technical Mem­ lowed by storage in 70% ethyl alcohol after 48 hours. orandum NOS NCCOS 27 (Kim et al., 2006). Five ran­ domly selected organisms were analyzed per site for all Shucking blue mussels usually results in severe damage histopathology measurements, except for Perkinsus mari- to the mantle lying next to the shell, so a more careful nus infection in oysters. For P. marinus, the Dermo disease approach was used to open their shells (Kim et al., 2006). pathogens in oysters, 12 oysters were analyzed per site. The tip of a sharp knife was carefully inserted between the shells at the ventral lip and run dorsally until the posterior While the actual quantification of parasites and disease adductor muscle was cut so that the shells remain in an is similar for all the bivalves, sample preparation differs open position. This was done to five mussels per site. Care for each bivalve species due to their diferent biology and was taken to cut no further than the adductor muscle to size. Brief descriptions of tissue sample preparation are avoid cutting into the digestive gland immediately below provided below. the adductor muscle. A sharp knife or scalpel was then carefully run between the shell and the mantle to sepa­ Bivalve Tissue Preparation rate the meat from the shell. This procedure was repeated for the second shell to completely detach both sides of For oysters, 12 were shucked with an oyster knife. From the mantle from the shell. The whole tissues were then each speciemen, a small section of mantle tissue (5 mm placed in Davidson’s fixative for one week and then trans­ x 5 mm) was removed for culturing of P. marinus patho­ ferred to 70% ethyl alcohol for storage. Once preserved, gen analysis. For other histopathology paramenters, a the tissue hardened and a 5-mm thick cross-section was 3–5 mm-thick transverse cross-section of tissue was re­ then removed using a scalpel. The cross-section was ob­ moved from five of the specimens using sharp scissors. tained such that the dorsal-ventral aspect passes through

5 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Parasites and Disease Quantification and graphing. Geographic information system ArcGIS® package was used for the spatial mapping of the results. Tissue sections were examined under the microscope

Methods using a 10× ocular and a 10× objective. When neces­ In this report, parasites or pathologies of the same taxa sary, a 25× or 40× objective was used for closer exam­ or group were pooled together by category, as indicated in ination. Tissue types examined include gill, mantle, go­ Tables 2a and 2b, and the resulting total values were used to nad and gonoducts, digestive gland tubules, stomach/ determine prevalence and intensity of parasites and disease. digestive gland, and connective tissue. Perkinsus marinus For instance, cestode parasites (tapeworms) include body in oysters was assayed by the more precise thioglycollate cestodes, gill cestodes, and mantle cestodes (Tables 2a). method (Powell and Ellis, 1998), rather than histology. All parasites and disease (Tables 2a and 2b) were count­ Prevalence describes the proportion of individ­ ed, either quantitatively or on a semi-quantitative scale. ual bivalves in a sampled population that are in­ fected by a specific parasite or carried a partic­ Conditions scored quantitatively were evaluated by ular pathology. Prevalence was calculated as: keeping a running count of number of parasites or dis­ eased tissues as slides were scanned to avoid re-exam­ ining each incident multiple times. Quantitative scores Prevalence= ( ( ∑ hosts with parasite or were used for parasites and disease that could be tallied pathology) / (number of hosts analyzed) ) x 100 individually (Kim et al., 2006). Parasites counted includ­ ed prokaryotic inclusion bodies (rickettsia and chlamydia), ciliates, gregarines, other protozoans, nematodes, encyst­ Intensity is a measure of severity of infection or disease ed cestodes, metacercariae of trematodes, copepods and in the affected bivalves. It was calculated as the average other unidentified organisms. Ciliate numbers were tal­ number of occurrences of the parasite or pathology in lied for each tissue type (gill and digestive tract), as were infected hosts. For conditons measured semi-quantitave­ gregarines. A number of tissue conditions were also eval­ ly, the scale rating replaced the number of occurrences. uated quantitatively, including ceroid bodies and hemo­ cytic infiltration, that were focal and diffuse in nature. Intensity = ( ∑ number of occurrences of parasite or pathology )/ (number of hosts with parasites or pathology) Some conditions were assigned to a semi-quantitative scale relative to the extent of infection or disease (Ta­ For each category of parasite and disease, prevalence and bles 3a - 3d). Definitions of scale values can be found in intensity were determined for each year on a site specific Kim et al. (2006). Haplosporidium nelsoni (MSX pathogen) basis. While the prevalence values varied from 0 (no-in­ infection in oysters was scored on a 0 to 6 point scale of fection or no-disease) to 100% (infection or disease), the Kim et al. (2006), adapted from Ford and Figueras (1988) intensity values varied broadly, and results were scaled (Table 3a). Perkinsus marinus infection in oyster mantle was from 0 to 100 using the formula: semi-quantified on a 0 to 5 point scale (Table 3b) estab­ lished by Craig et al. (1989). This was done on the mantle ((Site Value - Minimum Value) / (Maximum Value – Minimum tissue preparation instead of stained tissue sections. A 0 Value)) x 100 to 4 point scale was used for invasive trematode sporo­ cysts (Fellodistomidae and Bucephalidae) (Table 3c). For with maximum and minimum values for parasites digestive gland atrophy, the average degree of thinning and disease determined separately. of the digestive tubule walls was assigned a numerical rat­ ing on a 0 to 4 point scale (Kim et al., 2006) (Table 3d). Site-specific prevalence and intensity results based on the 2008 - 2009 Mussel Watch data, which repre­ sent the most recent and complete dataset, were used Data Analysis and Statistical Approach to evaluate spatial distributions of parasites and dis­ ease. A three-group scheme (low, medium, high) was All data processing and analysis were evaluated using applied to the site-specific prevalence and intensity val­ JMP® statistical software. Microsoft Excel and Sigma- ues using “Equal Interval” classification in ArcGIS. Plot® were also used for additional visual data analyses Thus, parasite infection or disease would be low, medi­

6 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters um or high for prevalence or intensity with values be­ The contaminants used in this assessment include major tween 0.0 - 33.3, 33.3 - 66.7, and 66.7 - 100, respectively. and trace metals, and the organic compounds as illustrat­ ed in Table 1. For this assessment, organic contaminants Spearman rank statistic was used to characterize re­ were grouped by compounds of the same class or by con­ lationships between contaminant body burden and geners. The concentration value of each class was defined parasite taxa and disease. When applicable, Wilcox- by calculating the sum (total) of the concentration of in­ Methods on or ANOVA were used to assess both region­ dividual compounds in the class. Thus, total PAHs was al and inter-species contrasts of parasitic infection. defined as the sum of 24 polycyclic aromatic hydrocarbon compounds; the sum of DDT and its metabolites, DDE To determine site-specific temporal trends, nonpara­ and DDD, as total DDTs; total dieldrins as the sum of metric Mann Kendall trend test was applied to the aldrin, dieldrin and lindane; the sum concentrations of long-term prevalence results for both parasite infec­ all chlordanes as total chlordanes; and the sum concen­ tion and disease. Because the MWP frequently estab­ trations of 18 individual PCB congeners as total PCBs. lished new monitoring sites, the “long term trends” were only assessed for sites with four or more years of data between 1995 and 2009. The Mann Kendall test assessed probability of consistent monotonic chang­ es of the prevalence values over the monitoring years.

Table 2a. List of parasites measured by MWP in sentinel bivalves.

Category Parasites Prokaryotes Digestive tubule rickettsia, Gut rickettsia, Chlamydia Gregarines Body gregarines, Gill gregarines, Mantle gregarines MSX H. nelsoni Dermo P. marinus Ciliates Digestive tract ciliates, Large gill ciliates, Small gill ciliates, Gut ciliates Cestodes Body cestodes, Gill cestodes, Mantle cestodes Trematodes Trematode sporocyst, Trematode metacercaria, Proctoeces Nematodes Nematodes Copepods Body copepods, Gill copepods, Gut copepods Pea crabs Pinnotherid crab

Table 2b. List of disease and host responses measured by the MWP in sentinel bivalves.

Category Disease Hemocytic infiltration Focal hemocytic infiltration, Diffuse hemocytic infiltration Ceroid bodies Ceroid bodies Digestive tubule conditions Digestive gland atrophy, Unusual digestive tubule Necrosis (tissue death) Focal necrosis, Diffuse necrosis Xenoma Xenoma

7 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

Table 3a. Semi-quantitative scale for Haplosporidium nelsoni (MSX) infection modified from Ford and Figueras

(1988). Methods Score Description 0 Uninfected, no parasites found in the tissue cross-section 1 Parasites confined to gill or digestive tract epithelial tissue, ≤ 10 plasmodia per 100X field of either gill or body tissue 2 Parasites restricted to gill or digestive tract epithelial tissue, Very light infection, 11 ≤ plasmodia ≤ 100 per 100X field of either gill or body tissue 3 Parasites spreading into gill or digestive tract subepithelium, parasites restricted to epithelium and subepithelium area, > 100 plasmodia per 100X field of either gill or body tissue but < 1 per 1000X oil immersion field 4 Parasites more evenly distributed in gill or digestive tract subepithelium and scattered through somatic tissue, > 100 per 100X field of either gill or body tissue but 1 to ≤ 10 per 1000X oil immersion field 5 Moderate systemic infection, averaging 11 to ≤ 20 parasites per 1000X oil immer­ sion field

6 heavy sistemic infection. averaging > 20 parasites per 1000X oil immersion field

Table 3b. Semi-quantitative scale for Perkinisus marinus [adapted from Mackin (1962) by Craig et al. (1989)].

Infection intensity Numeric value Descripytion Negative 0.00 No hypnospores present Very light 0.33 1-10 hypnospores Light 0.67 - 1.33 11 - >125 hypnospores but much less than 25% of tissue is hypnospores Light/moderate 1.67 - 2.33 <25% of tissue is hypnospores - >25% but much less than 50% of tissue is hypnospores Moderate 2.67 - 3.33 >25% but <50% of tissue is hypnospores - >50% but much less than 75% of tissue is hypnospores Moderately heavy 3.67 - 433 >50% but <75% of tissue is hypnospores - >75% but much less than 100% of tissue is hypnospores Heavy 4.67 - 5.00 >75% of tissue is hypnospores but some oyster tissue is still visible - Nearly 100% of tissue is hypnospores

8 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Table 3c. Semi-quantitative scale for trematode sporocyste.

Score Description

0 Uninfected Methods 1 Present in the gonads only (some gametic tissue still present) 2 Completely filling the gonads (no gametic tissue present); may be present in diges­ tive gland or gills in very limited amount 3 Completely filling the gonads, extensive invasion of the digestive gland and/or the gills 4 Completely filling the gonads; substantially filling the digestive gland or gill; indi­ viduals appear to be a sac of sporocyst

Table 3d. Semi-quantitative scale for digestive gland atrophy.

Score Description 0 Normal wall thickness in most tubules (0% atrophy), lumen nearly occluded, few tubules even slightly atrophied 1 Average wall thickness less than normal, but greater than one-half normal thick­ ness, most tubules showing some atrophy, some tubules still normal 2 Wall thickness averaging about one-half as thick as normal 3 Wall thickness less than one-half of normal, most tubules walls significantly atro­ phied, some walls extremely thin (fully atrophied) 4 Wall extremely thin (100% atrophied), nearly all tubules affected

9 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program RESULTS PARASITES Status and Temporal Trends Based on the 2008 - 2009 MWP monitoring data, the Bacteria (Prokaryotic Parasites) proportion of sentinel bivalves (excluding zebra mussels) parasitized by PIB was mostly low, as indicated by the prev­ Prokaryotic Inclusion Bodies alence values (Figure 3). The infection rates were most­ (Chlamydia-like and Rickettsia-like) ly lower at blue mussel sites. At oyster sites, prevalence ranged from low to medium, except at locations such as Overview Cape Henlopen, DE and Cape May, NJ where high preva­ lence values were found (Figure 3). PIB infection intensity Chlamydia and rickettsia are included together under was generally low even at locations where medium to high Results Parasites Results the heading of prokaryotic inclusion bodies (PIB) (Fig­ prevalence values were found among oysters (Figure 4). ure 2). Chlamydia are non-motile, coccoid bacteria that replicate within the cytoplasmic vacuole in the host. Over the monitoring assessed in this study, PIB infec­ They have two forms, a reticulate and an elementa­ tion was found to be fairly static at most of the mon­ ry body, which are infectious particles (spore-like) that itoring sites, except at a few sites where significant (p are released when the infected cell ruptures. Rickett­ < 0.05) trends were detected (Figure 5). A decreas­ sia are also non-motile intracellular parasites. They can ing trend (p = 0.007) was observed at a site in Mission replicate either within the nucleus or cytoplasmic vac­ Bay, CA, while a couple of notable increasing trends uole of the host. Several rickettsial bacteria have been were found at monitoring sites in Palmico Sound, linked to human diseases, such as typhus, rickettsial pox NC (p = 0.04) and Sapelo Sound, GA (p = 0.03). and Rocky Mountain spotted fever. PIBs are usually ob­ served in the duct and tubule walls of the digestive gland Correlation of oysters and blue mussels (Kim et al., 2006). As a part No significant correlations were found between con­ of the MWP, PIBs are not measured in zebra mussels. taminant body burden and prevalence of PIB infection.

Figure 2. Prokaryotic inclusion bodies (PIB) in digestive tract epithelium of an American oyster. Arrows indicate PIBs (Kim et al., 2006).

p.10 10 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 3. Spatial distribution of prokaryotic inclusion bod­ ies (chlamydia and rickettsia) prevalence in oysters and blue mussels, based on 2008 - 2009

Mussel Watch data. Results Parasites Results

Figure 4. Spatial distribution of prokaryotic inclusion bodies (chlamydia and rickettsia) in­ fection intensity in oysters and blue mussels, based on 2008 - 2009 Mussel Watch data.

Figure 5. Site-specific long- term temporal trends of prokaryotic inclusion bodies (chlamydia and rickettsia) prevalence in oysters and blue mussels.

p.11 11 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Protozoan Parasites (Gregarines) southern California, and in oysters from one site in south­ ern Puerto Rico. These findings corroborate results by Overview Kim and Powell (2007), who found that gregarines are dominant parasites in oysters and are distributed along the Gregarine parasites require two hosts; a mollusk for entire range of the East Coast and Gulf of Mexico. How­ the intermediate stage (Figure 6), and an arthropod as ever, the severity of the infection was mostly low with only the final host (Meyers and Burton, 2009). They­ usual a high infection intensity found in Apalachicola Bay, FL, ly invade the intestines, but are also found in other tis­ and mediun intensity found in Pass Christian, MS, Co- sues, such as the foot, mantle, and palps. Kim and pus Christi, Matagorday Bay, and Mesquite, TX (Figure 8). Powell (2007) found gregarines in the connective tissue Gregarine infection was static at most of the monitoring Results Parasites Results surrounding the visceral mass, as well as in the gills and the mantle connective tissues of blue mussels and oys­ sites, except at few sites where significant trends (p < 0.05) ters. Although heavy infections have been suggested to were observed (Figure 9). While sites in Delaware Bay, NJ have some harmful effects on oyster physiology (Sind­ and Florida Bay, FL showed decreasing trends, gregarine ermann, 1990), Cheng (1967) concluded that, in general, infection was found to be on the rise at locations in: Mon­ gregarines infections have low pathogenicity in bivalves. terey Bay and Santa Monica Bay, CA; San Antonio Bay According to Meyers and Burton (2009), the presence and Montagorda Bay, TX; Lake Borgne, LA; Florida Bay, of gregarines in bivalves causes no human health con­ FL; Chesapeake Bay, VA; and Delaware Bay, NJ (Figure 9). cerns. Gregarines do not typically occur in zebra mussels. Correlation Status and Temporal Trends In blue mussels from the West Coast, gregarine Gregarines were found in the American oyster and blue prevalence was positively correlated with arse­ mussels at variable prevalences (Figure 7). Oysters from nic and lead, but negatively correlated with man­ the Gulf of Mexico and the eastern seaboard, from Flor­ ganese body burden (p < 0.001) (Appendix A). ida to Virginia, had the highest infection prevalence (Fig­ ure 7). Elevated prevalence of gregarines was also found in blue mussels from coastal waters of Washington and

Figure 6. Section of oocyst (arrow) of gregarine Nematopsis sp. in the connective tissues of scallop (left); in oyster (right) (Meyers and Burton, 2009).

p.12 12 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 7. Spatial distribution of gregarine prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch

data. Results Parasites Results

Figure 8. Spatial distribution of gregarine infection intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data.

Figure 9. Site-specific long- term temporal trends of gregarine prevalence in oysters and mussels.

p.13 13 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Haplosporidium nelsoni (MSX Pathogen) Status and Temporal Trends

Overview Prevalence and intensity results indicate that H. nelsoni was present in oysters, but at low prevalence and intensity Haplosporidium nelsoni is a protozoan that is the etiolog­ (Figures 11 and 12). Even in Chaseapeake Bay, where his­ ical agent of multinucleated sphere X/unknown (MSX) torical outbreaks of the parasites have been reported (An­ disease in oysters (Figure 10). MSX infections start in the drews, 1979; Kemp et al., 2005), the infection was not se­ gill epithelium, and can remain at light infection levels for vere (Figure 12). The Chesapeake Bay Program considers months before the disease worsens and becomes systemic MSX as among the major biological stressors that cause (Ewart and Ford, 1993). Oysters with advanced infection the most oyster mortality (Chesapeake Bay Program, on-

show mantle recession, gaping valves, watery emaciated line). Since 1996, the Maryland Department of Natural Results Parasites Results tissues, a pale digestive gland, and often begin to die with­ Resouces (MD DNR) has conducted an annual survey in a month. Death of these oysters is so rapid that there to assess the status of oysters in the Chesapeake Bay. is no loss of soft tissue (Haskin et al., 1965; Andrews, Our results at the time of the study were in agreement 1966; Couch and Rosenfield, 1968). MSX disease, which with MD DNR, which reported an average prevalence was reported in Chesapeake Bay in the 1950s, was a ma­ value of 30% for H. nelsoni in 2008 (Tarnowski, 2010). jor contributor to the devastation of oyster fisheries in the Chesapeake Bay (Andrews, 1979), and along the East Temporal variation of H. nelsoni infection prevalence Coast in the 1980s (Kemp et al., 2005). Although the par­ was found to be static (p > 0.05) at all oyster sites on the asite is lethal to oysters, it is said to be harmless to humans East Coast (Figure 12). This was an indication that H. nel- (Ewart and Ford, 1993). As part of the MWP, H. nelsoni is soni infection, hence MSX disease, remained fairly stable only measured in East Coast oysters. in these oysters over the monitoring period (Figure 13).

Correlation

The data showed no correlation between H. nelsoni and the measured contaminants in the oysters examined.

Figure 10. Numerous multinucleated plasmodia of Haplosporidium nelsoni in the gills of an American oyster. Arrows point to example parasites (Kim et al., 2006).

p.14 14 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 11. Spatial distribution of H. nelsoni prevalence in oysters, based on 2008 - 2009

Mussel Watch data. Results Parasites Results

Figure 12. Spatial distribution of H. nelsoni infection intensity in oysters, based on 2008 - 2009 Mussel Watch data.

Figure 13. Site-specific long- term temporal trends of H. nelsoni prevalence in oysters.

p.15 15 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

Perkinsus marinus (Dermo Pathogen) coast (Grout, 2011). The decline of the oyster population in the Chesapeake Bay has been attributed to many fac­ Overview tors, including Dermo disease and mortality (Tarnowski, 1999). Perkinsus marinusis a protozoan that is the etiological agent of Dermo (Perkinsosis), a major disease that causes Status and Temporal Trends high mortality in oysters (Figure 14). Perkinsus marinus can be transmitted directly from oyster to oyster and occurs The parasite is highly prevalent in both Gulf Coast through the water column during feeding (Elston, 1990). and East Coast waters (Figure 15). The distribution of Proliferation of the parasite within the host causes sys­ P. marinus prevalence mirrored the historic description of

temic disruption of connective tissue and epithelial cells, the Dermo pathogen along the Atlantic seaboard and the Results Parasites Results and is correlated with warm summer water temperatures Gulf of Mexico (ICES, 2012). For their 2008 survey, the (higher than 20 °C and higher salinity), when pathogenic­ MD DNR reported similar prevalence values for P. mari- ity and associated mortalities are highest (ICES, 2012). nus infection in oyster from the Chasepeake Bay, with sev­ Oyster death is usually a consequence of rapid prolifer­ eral sites having more than 70% prevalence (Tarnowski ation of the parasites invading the oyster, lysing tissues, 2010). However, infection intensity was low at all the oyster and occluding hemolymph vessels. Kennedy et al. (1996) sites, including sites from the Chesapeake Bay (Figure 16). explained that P. marinus stunts oysters’ ability to produce new shell deposits and impairs adductor muscle strength, Temporal variation of P. marinus infection prevalence was which leaves the host weak and prone to gape (dead but found to be static (p > 0.05) at all oyster sites (Figure 17). with soft tissue still present) when removed from water. This was an indication that P. marinus infection, hence Der- American oysters tend to be most susceptible to infec­ mo disease, remained fairly stable in oysters from the Gulf tion, although clams and other mollusks are susceptible of Mexico and East Coast over the monitoring period. to infection by various Perkinsus species. The parasites have not been associated with disease in mussels. Histor­ Correlations ical outbreaks of Dermo disease have been linked to high No significant correlations were found between contam­ oyster mortality along the southern and middle Atlantic inant body burden and prevalence of P. marinus infection.

Figure 14. Several Perkinsus marinus cells in the intestine of an American oyster (40x mag, MHE stained). Photo by Joe Marcino, Maryland Department of Natural Resources. p.16 16 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 15. Spatial distribution of P. marinus prevalence in oysters, based on 2008 - 2009

Mussel Watch data. Results Parasites Results

Figure 16. Spatial distribution of P. marinus infection inten­ sity in oysters, based on 2008 - 2009 Mussel Watch data.

Figure 17. Site-specific long- term temporal trends of P. marinus prevalence in oysters.

p.17 17 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Ciliates Status and Temporal Trends

Overview The prevalence of ciliate infections was low to medi­ um at the vast majority of both mussel and oyster sites Ciliates are single-celled eukaryotes with hair-like or­ (Figure 19). However, hot-spots for ciliate infection were ganelles called cilia, which are used for movement and found with site-specific prevalence values that are more food collection (Figure 18). Ciliates can be found in every than 70%, although the intensity of the infections is gen­ aquatic system, such as ponds, lakes, rivers, and oceans. erally low (Figure 20). They reproduce primarily by cell division. However, in some instances, two cells can fuse together to form new Ciliate infection was static, with no significant increasing organisms. Parasitic ciliates are transmitted from host to or decreasing trends (p > 0.05) over the monitoring peri­ Results Parasites Results host through the water column and can cause serious tis­ od (Figure 21). sue damage or stress in the host organisms (Meyers and Burton, 2009). Ciliates are found in the gills, gut, and di­ Correlations gestive tract of oysters and blue mussels, but not in zebra Ciliate prevalence was positively correlated with manga­ mussels (Kim et al., 2006). Ciliate infections do not appear nese in blue mussels from the West Coast (Appendix A). to elicit obvious pathological conditions or host responses in blue mussels and oysters, however, some mature and multiplying ciliates can cause infected cells to distend and form a xenoma (Brandão et al., 2013). Xenoma is present­ ed later in this report.

Figure 18. Ciliates in the intestine of an oyster (left) and between gill filaments of a blue mussel (right) (Kim et al., 2006).

p.18 18 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 19. Spatial distribution of ciliate prevalence in oysters and mussels, based on 2008 -

2009 Mussel Watch data. Results Parasites Results

Figure 20. Spatial distribution of ciliate infection intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data.

Figure 21. Site-specific long- term temporal trends of ciliate prevalence in oysters and mussels.

p.19 19 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

Metazoan Parasites Status and Temporal Trends The spatial distribution based on prevalence values (Fig­ Cestodes (Tapeworms) ure 23) and infection intensity (Figure 24) indicate that infection by cestodes, which were only recorded in oys­ Overview ters, were generally low. However, some hot-spots were Cestodes are a class of parasitic flatworms commonly observed at a few sites, with prevalence values ranging called tapeworms due to the extended length of the adults. from medium to high. In the Gulf of Mexico, hot-spots Cestodes are hermaphrodites, but self-fertilization is rare. for cestode infection included Apalachicola Bay and Cross-fertilization is the main reproduction practice, thus Tampa Bay, FL and Lower Laguna Madre, TX. Along

Results Parasites Results additional organisms are essential for the continuation of the East Coast, these hot-spots were also found in Alta­ the species within the host. The life cycle of tapeworms maha River, GA, and Beaufort Inlet, NC. Cestode infec­ requires one intermediate host (e.g. bivalves), which har­ tion intensity was low in all infected oysters (Figure 24). bor eggs and larvae (encysted sporocysts and metacer­ Cestode infection prevalence in oysters was found to be cariea larvae), and one definitive host (vertebrates, in­ static (p > 0.05). This was an indication that, although some cluding humans), which harbor the adult worms (Roberts hot-spots were observed, the infection rate remained fair­ et al., 2005). There are over a thousand species of tape­ ly stable troughought the monitoring period (Figure 25). worms that parasitize various animals, including oysters, but they are not found in blue mussels and zebra mussels. Correlations Transmission to humans can occur through consumption of raw shellfish and fish. In humans, light infections are Cestode infections in American oysters were sig­ usually non-symptomatic. However, abdominal discom­ nificant, and correlated (p < 0.0001) with trace fort, gastric pain followed by vomiting and diarrhea can metal body burdens (Appendix A), with a posi­ occur in heavier infections (Bogitsh and Carter, 2005). In tive association with arsenic, and negative correla­ bivalves, host response to the infection is characterized tions with both cadmium and nickel body burdens. by encapsulation of larval cestodes in connective tissue (Figure 22). Thus, the infection does not seem to signifi­ cantly cause harm to the host bivalves (Cheng, 1966).

Figure 22. Encapsulated cestode larvae in the vesicular connective tissue surrounding the stomach of an American oyster (Kim et al., 2006).

p.20 20 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 23. Spatial distribu­ tion of cestode prevalence in oysters, based on 2008 - 2009

Mussel Watch data. Results Parasites Results

Figure 24. Spatial distribution of cestode infection intensity in oysters, based on 2008 - 2009 Mussel Watch data.

Figure 25. Site-specific long- term temporal trends of cesto­ de prevalence in oysters.

p.21 21 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

Trematodes (Flatworms, Flukes) Status and Temporal Trends

Overview Trematodes can infect both mussels and oysters, but in­ fection distribution based on the 2008 - 2009 MWP data Trematodes are a group of parasitic flatworms within shows a clear regional pattern, with low prevalence ob­ the phylum Platyhelminthes, that have a distinctive exter­ served at all the monitoring sites, except at sites in the nal feature of having two suckers, one near the mouth and Northeast region (Figure 27). Trematode infection preva­ the other on the organism’s ventral surface. Figure 26 de­ lence ranged mostly from medium to high at blue mussel picts a Bucephalus sp. trematode infection in a blue mussel sites along the Northeast corridor. Trematode infection (Meyers and Burton, 2009). Nearly all trematodes are par­ intensity was low at all infected sites (Figure 28), howev­

asites of freshwater and marine mollusks and vertebrates, er, high prevalence values in the Northeast suggest that Results Parasites Results and require two host species to complete their life cycle. the blue mussels there are more suceptible to trematode As larvae, they live in the digestive and reproductive tis­ infection. sues of bivalves, often causing sterilization (Cheng, 1967; Sindermann, 1990). If the parasite is present in large num­ Trematodes infection was mostly static, however, signif­ bers, it can cause tissue destruction and death (Meyers and icant decreasing tempral trends were found at two sites Burton, 2009). Meyers and Burton (2009) also observed (Figure 29). Trematode infection was found to be decreas­ that, in mussels, trematodes can lower byssal thread pro­ ing at the blue mussel site of Hingham Bay in Boston duction, cause infertility, and affect pearl formation. If in­ Harbor, MA (p = 0.03), and at the oyster site at Mosquito gested by humans, trematodes can cause severe intestinal Point in San Antonio Bay, TX (p = 0.04). illnesses (Meyers and Burton, 2009). Trematodes do not Correlations typically occur in zebra mussels. No significant correlations were found between con­ taminant body burdens and trematode infection.

Figure 26. Trematode sporocyst (Bucephalus sp.) infection in a blue mussel (Meyers and Burton, 2009).

p.22 22 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 27. Spatial distribution of trematode prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch

data. Results Parasites Results

Figure 28. Spatial distribution of trematode infection inten­ sity in oysters and mussels, based on 2008 - 2009 Mussel Watch data.

Figure 29. Site-specific long- term temporal trends of trem­ atode prevalence in oysters and mussels.

p.23 23 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Nematodes do not typically occur in blue mussels. Nematodes (Roundworms) Status and Temporal Trends Overview Netmatodes are measured in zebra mussels and oys­ Nematodes (roundworms) are the second most diverse ters, which are most likely to harbor larval stages of group of animals on the planet, second only to arthropods. the parasites (Kim et al., 2006). The data indicated that Roundworms have a tubular digestive system with open­ nematode infection was very low, as illustrated by the ings at both ends of a cylindrical body. Over half of the prevalence and intensity values (Figures 31 and 32). 28,000 described species are parasitic (Hugot et al., 2001). However, medium range infection prevalence values They have multiple development stages (Cheng, 1978). were found at a few sites in the Gulf of Mexico, in­ Results Parasites Results Cheng (1966) suggested that larval nematodes invade cluding Matagorda Bay, TX; Breton Sound, LA; Flor­ oysters via the digestive tract and migrate through tissues ida Bay, FL; and Mississippi Sound, MS (Figure 31). by way of blood vessels. In some cases, host responses include infiltration of hemocytes around the area where Nematode infection prevalence was found to be stat­ the worm is located (Kim et al., 2006). Infections in mol­ ic (p > 0.05) at all sites. This indicated that the infec­ lusks can cause destruction of adjacent host tissue as well tion of the worms in oysters and zebra mussels has been as the formation of granulomas in infected cells (Mey­ fairly stable over the monitoring period (Figure 33). ers and Burton, 2009). Nematodes that infect mollusks, such as oysters and zebra mussels, are mainly larval stages Correlations (Figure 30), while adult nematodes are found in predators of mollusks. Adult forms often live in the digestive tract A positive (p < 0.001) correlation was observed be­ of vertebrates, like humans, where they can cause serious tween nematode infection prevalence and cadmi­ gastrointestinal health issues (Bogitsh and Carter, 2005). um body burden in American oysters (Appendix A).

Figure 30. Unidentified nematode larvae in connective tissue of the digestive gland of an American oyster (Kim et al., 2006).

p.24 24 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 31. Spatial distribution of nematode prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch

data. Results Parasites Results

Figure 32. Spatial distribution of nematode infection inten­ sity in oysters and mussels, based on 2008 - 2009 Mussel Watch data.

Figure 33. Site-specific long- term temporal trends of nem­ atode prevalence in oysters and mussels.

p.25 25 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Copepods Status and Temporal Trends

Overview Copepod infection was found to be generally low in all sentinel bivalves, as indicated by the prevalence and Copepods are small animals of the phylum Arthrop­ intensity values (Figures 35 and 36). Medium range oda, subphylum Crustacea. They are aquatic animals prevalence values were found at blue mussel sites, in­ with exoskeletons. Parasitic copepods lay their eggs in cluding Santa Monica Bay and Tijuana River estuary, the water and are subsequently ingested by filter-feeding CA; Possession Point, WA; and Jamaica Bay, NY. Oys­ bivalves (Heegaard, 1962; Darwin and Stefanich, 1966). ters from Lake Barre in LA were also found to harbor They are mainly found in gills (Figure 34), but can also be the parasite at medium infection prevalence (Figure 35). found in the digestive tract. Appendages of the copepods Results Parasites Results (e.g. antennae) can cause erosion and metaplasia of the Copepod infection was found to be static (p > intestinal epithelium. It is estimated that nearly half of 0.05) at all sites, indicating infection rates were the 13,000 species of copepods are parasitic (Heegaard, stable over the monitoring period (Figure 37). 1962). Parasitic copepods infecting bivalves may be ei­ ther obligate endoparasites, which affect the alimentary Correlation tract, or ectoparasites, which affect the mantle and gills. No significant correlations were found between contaminant body burdens and copepod infection.

Figure 34. Copepod (*) in gill surrounded by an intense hemocyte reaction (Carballal et al., 2001).

p.26 26 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 35. Spatial distribu­ tion of copepod prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch

data. Results Parasites Results

Figure 36. Spatial distribution of copepod infection intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data.

Figure 37. Site-specific long- term temporal trends of cope- pod prevalence in oysters and mussels.

p.27 27 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Pinnotherid Crabs (Pea Crabs) Status and Temporal Trends

Overview Pea crab infections were at low prevalence and in­ tensity at virtually all blue mussels and oyster sites na­ Pea crabs are small crustaceans in the Pinnotheridae tionwide, with the exception of some locations along family, which exclusively parasitizes bivalves (Figure 38). the Atlantic and Gulf coasts, where medium infec­ Pea crabs are occasionally found in the mantle cavity of tions were observed (Figures 39 and 40). The most se­ oysters and blue mussels, but not in zebra mussels (Kim et vere pea crab infections were found at Dandy Point in al., 2006). Small pea crabs can also be found in the gills or Chesapeake Bay, VA; Altamaha River, GA; and Port in the water-conducting channels in the back of gills (Ha­ Isabel and South Bay in Lower Laguna Madre, TX. ven et al., 1978). Pea crabs can have detrimental impacts Results Parasites Results on the host by removing food particles captured in the Pea crab infection temporal trend was found to be stat­ bivalve’s gill, which over time can lead to food deprivation ic (p > 0.05) at all sites (Figure 41). This indicated that (Kennedy et al., 1996). Also, direct injury can occur when infection has been stable over the monitoring period. the crab is connected to the host’s gill by causing erosion, and the walking legs damage the gill tissue as it seeks food Correlation in the mucous strings (Stauber, 1945; Meyers and Burton, No significant correlations were found between 2009). Bivalves with pea crabs have been found to contain contaminant body burdens and pea crab infection. less tissue mass per shell cavity volume (Haven, 1958).

Figure 38. A pea crab (Pinnotheridae family) living inside an American oyster (NOAA Ocean Explorer).

p.28 28 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 39. Spatial distribution of pinnotherid crab prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch

data. Results Parasites Results

Figure 40. Spatial distribution of pinnotherid crab infection intensity in oysters and mus­ sels, based on 2008 - 2009 Mussel Watch data.

Figure 41. Site-specific long- term temporal trends of pinnotherid crab prevalence in oysters and mussels.

p.29 29 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

Results DISEASE Status and Temporal Trends Hemocytic infiltration condition was observed at vari­ Tissue Inflammation able prevalence in all sentinel bivalves. The spatial dis­ (Tissue Hemocytic Infiltration) tribution showed the presence of high prevalence sites along the East and Gulf coasts (Figure 43). Along the Overview West Coast, the condition occured mostly at low prev­ alence, except for a few sites with medium range prev­ Tissue inflammation is characterized by the infiltration

Results Disease Results alence (Figrue 43). The condition had generallly low of hemocytes (Figure 42) in response to foreign bodies, prevalence in the Great Lakes, Alaska, Hawaii and Puer­ parasites, and other stressors. Bivalve hemocytes move to Rico, but in Bear Cove, Cook Inlet, AK; and Ba­ freely amongst all tissues and are considered the primary hia de Boqueron, PR; medium prevalence values were cellular defense mechanism (Soudant et al., 2013). Hemo­ found (Figure 43). The intensity of hemocytic infiltra­ cytic infiltration is a response or defense mechanism to tion condition was low in all the bivalves (Figure 44). injury or invasion of non-host material. The condition is also called “inflammatory response.” Hemocytes phago­ Hemocytic infiltration was found to be fairly sta­ cytize or engulf non-host material in infected tissue and ble at most of the monitoring sites. However, the dis­ attempt to remove them from the organism. Their pur­ ease was found to be increasing (p < 0.05) at some pose is to neutralize or isolate potentially damaging sub­ sites (Figure 54). These include sites in Boston Har­ stances (Kennedy et al., 1996). Hemocytic infiltration bor, MA; Quinby Inlet, VA; Brazox River, TX; Monte­ may be focal or diffuse. Diffuse is distinguished from fo­ rey Bay and Palos Verdes, CA; and Puget Sound WA. cal when hemocytes are distributed broadly over a large section of tissue without a clear center or focal point Correlations of highest hemocyte concentration (Kim et al., 2006). No significant correlations were found between con­ taminant body burdens and hemocytic infiltration.

Figure 42. Hemocytic infiltration (arrow) near the gill base of an American oyster (Kim et al., 2006).

30 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 43. Spatial distribution of hemocytic infiltration prev­ alence in oysters and mussels, based on 2008 - 2009 Mussel

Watch data. Results Disease Results

Figure 44. Spatial distribution of hemocytic infiltration inten­ sity in oysters and mussels, based on 2008 - 2009 Mussel Watch data.

Figure 45. Site-specific long- term temporal trends of tissue hemocytic infiltration preva­ lence in oysters and mussels.

31 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

Ceroid Bodies found in American oysters (Figure 47); this is consistent with previously published results (Kim et al., 2006). Even Overview though ceriod bodies were highly prevalent, the intensity of the disease was mostly low (Figure 48). At a few oyster Ceroid is a form of lipofuscinosis, a metabolic cellu­ sites, including Charlotte Harbor, St. Johns River, Matan­ lar disease (Figure 46). It is caused by a lack of enzymes, zas River, and Cedar Key, FL; Atchafalaya Bay, LA; Swan which allows waste products (ceroid bodies or lipofus­ Point, MD; and Mesquite Bay and Nueces Bay, TX; ceroid cin) to accumulate in cells (Zaroogian and Yevich, 1993). intensity was medium, but the most severe case of ceroid

Results Disease Results Ceroid bodies are brownish-yellow aggregates linked to disease was found in Charlotte Harbor, FL (Figure 48). metabolite accumulation and detoxification (Zaroogian and Yevich, 1993). Ceroids can interfere with normal cell The temporal trend of ceroid prevalence was found to functioning, and are also linked to cell aging in bivalves. be static (p > 0.05), indicating that the disease has been relatively stable over the monitoring period (Figure 49). Status and Temporal Trends Correlations Prevalence and intensity results show a broad distribu­ tion of ceroid disease in U.S. coastal waters (Figures 47 Prevalence of ceroid bodies was positively correlat­ and 48). High ceroid prevalence values were found in blue ed with arsenic body burdens in blue mussels from mussels and oysters along the East and West Coasts, as well the West Coast (Appendix A). Although a positive as in the Gulf of Mexico. High prevalences of the disease correlation does not imply direct effects, in regard were also observed in Bahia de Boqueron and Bahia Mon­ to chemical contaminants, positive correlations may talva, PR (Figure 47). In general, ceroid bodies were more suggest a cellular response as a result of cell stress. frequent in oysters, with 81% of the high prevalence cases

Figure 46. Ceroid bodies in gonad tissue of an American oyster (40x mag, MHE stained). Photo by Joe Marcino, Maryland Department of Natural Resources.

32 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 47. Spatial distribution of ceroid body prevalence in oysters and mussels, based on

2008 - 2009 Mussel Watch data Results Disease Results

Figure 48. Spatial distribution of ceroid body intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data.

Figure 49. Site-specific long- term temporal trends of ceroid body prevalence in oysters and mussels.

33 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

Digestive Gland Atrophy Status and Temporal Trends

Overview Of all the histopathology parameters measured, at­ rophy of the digestive gland was the most common Atrophy of the digestive gland (Figure 50) is character­ and widespread. High prevalence levels were found ized by thinning of the tubule walls (Winstead, 1995), that in all species measured and all locations, except the can lead to poor nutrition, poor growth and eventually western Great Lakes (Figure 51). Although the dis­ to low reproduction in bivalves. It is linked to stressors, ease was highly prevalent, it was not severe at any of

Results Disease Results such as exposure to contaminants and poor nutrition the monitoring sites, as intensity was low (Figure 52). (Winstead, 1995). Atrophy of the digestive gland has been used, along with biomarkers, to evaluate the environ­ The prevalence of the digestive gland atrophy prev­ ment’s recovery after an oil spill (Garmendia et al., 2011). alence was static (p > 0.05) at all monitoring sites (Fig­ Kim and Powell (2009) have also made linkages of trends ure 53). This indicated that although the condition is of digestive tubule atrophy to interannual variation in cli­ prevalent, it has remained stable over the monitoring mate, such as El Niño. period (Figure 53).

Correlation

No significant correlations were found between contaminant body burdens and prevalence values of digestive gland atrophy.

Figure 50. Atrophy (red arrow) of the digestive gland in an American oyster characterized by atrophic epi­ thelia resulting in rounded and enlarged lumina compared to quadriradiate lumina (black arrow) formed by normal thick epithelium (Kim et al., 2006).

34 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 51. Spatial distribution of digestive gland atrophy prevalence in oysters and mussels, based on 2008 - 2009

Mussel Watch data. Results Disease Results

Figure 52. Spatial distribution of digestive gland atrophy in­ tensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data.

Figure 53. Site-specific long- term temporal trends of diges­ tive gland atrophy prevalence in oysters and mussels.

35 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

Tissue Necrosis (Tissue Death) Status and Temporal Trends

Overview Prevalence and intensity of tissue necrosis were very low (Figures 55 and 56). A monitoring site located at the Los Necrosis (Figure 54) is an advanced stage of tissue Angeles River mouth, CA was the only monitoring site breakdown that is distinguished from other forms of with medium tissue necrosis prevalence (Figure 55). Low tissue death by drastic changes visible to the naked eye intensity values of the disease were only found at sites in (Majno and Joris, 1995). It is generally caused by factors southern California and the Northeast Atlantic (Figure 56).

Results Disease Results external to the host’s living cells or tissues. In bivalves, necrosis is often caused by pathogen infection (Meyers Temporal trends of tissue necrosis prevalence and Burton, 2009). Necrosis does not elicit the immune was static (p > 0.05) at all monitoring sites (Fig­ system by sending chemical signals, so necrotic cells are ure 57). This indicated that the disease was fair­ not removed by phagocytes and build up within the bi­ ly stable during the monitoring period (Figure 57). valve tissue. Bacillary necrosis disease in larval and juve­ nile bivalves often results in extensive cellular destruc­ Correlation tion that causes high mortality (Tubiash et al., 1965). No significant correlations were found between contami­ nant body burdens and prevalence values of tissue necrosis.

Figure 54. Necrosis in the intestine of an American oyster (20x mag, MHE stain). Cells with pyknotic nuclei (arrow). Photo by Joe Marcino, Maryland Department of Natural Resources.

36 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 55. Spatial distribution of tissue necrosis prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch

data. Results Disease Results

Figure 56. Spatial distribution of tissue necrosis intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data.

Figure 57. Site-specific long- term temporal trends of tissue necrosis prevalence in oysters and mussels.

37 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

Xenomas Status and Temporal Trends

Overview Overall, the prevalence of xenoma was low (Figure 59). Among blue mussels, there was only a single me­ Xenoma (Figure 58) is defined as an enlargement of dium prevalence site, Elliot Bay in Puget Sound, WA. tissue infected by parasites (Kim et al., 2006). When Among oyster sites, Lake Charles in Calcasieu Lake, LA intracellular parasites accumulate within host cells, it and Mullet Key Bayou in Tampa Bay, FL also displayed causes hypertrophy (xenoma) of the infected cell and medium prevalence values. The intensity of the disease

Results Disease Results its nucleus (Boehs et al., 2009). Multiplying ciliates are was equally low (Figure 60). Sites with low xenoma inten­ known to cause xenoma in oysters and blue mussels. sity were found mostly in the Gulf of Mexico, but also Xenoma causes local epithelial erosion and most like­ in Elliot Bay and Edmonds Ferry in Puget Sound, WA. ly impedes internal water flow (Scarpa et al., 2006). The condition does not typically occur in zebra mussels. Temporal trend of xenoma prevalence was stat­ ic (p > 0.05) at all oyster and blue mussels sites (Fig­ ure 61). This indicated that the disease was fairly sta­ ble throughout the monitoring period (Figure 61).

Correlation

No significant correlations were found between con­ taminant body burdens and prevalence values of xenoma.

Figure 58. Xenomas filled with ciliates in gills of the mangrove oyster in early (left) and more advanced stages (right). Arrows = ciliates; arrowheads = nucleus of the host cell (Boehs et al., 2009).

38 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Figure 59. Spatial distribu­ tion of xenoma prevalence in oysters and mussels, based on 2008 - 2009 Mussel Watch

data. Results Disease Results

Figure 60. Spatial distribution of xenoma intensity in oysters and mussels, based on 2008 - 2009 Mussel Watch data.

Figure 61. Site-specific long- term temporal trends of xenoma prevalence in oysters and mussels.

39 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

SYNTHESIS marinus average prevalence values, which were similar to survey results reported by the MD DNR in Chesapeake Parasites and disease are broadly distributed in oysters Bay. and mussels at the national scale. On average, most of the histopathology parameters have mostly low (< 33.3%) to Additionally, for parasites and disease, the proportion

Synthesis medium (33.3% - 66.6%) prevalence or intensity values, of bivalves that were infected was compared among the indicating a fairly minor infection rate in U.S. coastal wa­ bivalve types (Table 6), and between regions (Table 7). ters. This may be due to the fact parasite infection and Prevalence of ciliates, pinnotherid crabs, xenoma, and di­ diesease vary seasaonally with fluctuating temperature, gestive tubule atrophy did not vary significantly among and since the NS&T MWP only collect bivalve samples the bivalve species (p > 0.05). Prevalence and intensity for analysis in the winter, the optimum infection peri­ of all other parasite taxa and disease varied (p < 0.05). od of the year may have been missed. However, para­ In general, oysters had significantly higher prevalence of site infection and disease hot-spots were also observed parasite infections and disease than mussels (p < 0.05). throughout, with high prevalence values. It is noteworthy However, East Coast blue mussels had significantly higher to highlight the fact that digestive gland atrophy, which prevalence (p < 0.05) of tissue necrosis, digestive tubule can impact nutrition, and subsequently growth and re­ atrophy, and trematodes infection. While zebra mussels production, is highly prevalent in all bivalves and in all harbored fewer parasites and significantly lower cases coastal areas. The average prevalence and intensity values of disease, blue mussels had medium to high prevalence for each histopathology parameter are summarized by bi­ of parasitic infections. Dreissena polymorpha was first dis­ valve species and coastal region (Tables 4 and 5). Where covered in the Great Lakes in 1988 (Hebert et al., 1989). available, the MWP 2008 - 2009 data compared well with Quagga mussels (Dreissena bugensis) were found in 1989 some regional data, as in the case of H. nelsoni and P. and named a distinct species in 1991 (May and Marsden,

Table 4. Average prevalence (%) of parasites and diseases by region and by bivalve type (nd indicates that pa­ rameter was not detected in a given bivalve species) based on the 2008 - 2009 MWP data.

Parameter American American Blue mussel Blue mussel Zebra oyster (Gulf) oyster (East) (East) (West) mussel (GL) Prokaryotes 5.5 12.7 8.4 1.1 0.0 Gregarines 78.5 70.7 1.3 34.5 nd H. nelsoni nd 4.0 - - - P. marinus 46.9 41.4 - - - Ciliates 15.7 28.0 28.4 11.6 nd Cestodes 13.5 18.7 nd nd nd Trematodes 1.5 0.0 47.6 0.9 nd Nematodes 10.2 2.0 nd nd 0.0 Copepods 2.5 1.3 3.1 4.3 0.0 Pea crabs 1.5 4.7 0.9 0.0 - Hemocytic infil­ 19.7 12.7 33.8 12.3 0.9 tration Ceroid bodies 81.2 86.0 21.3 29.5 4.5 Digestive tubule 88.9 92.0 100.0 99.8 87.3 conditions Necrosis 0.0 0.0 4.4 1.6 0.0 Xenoma 4.6 0.0 0.0 0.9 nd “-” indicates that the histopathological parameter does not occur in the organism.

40 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Table 5. Average intensity of parasite infections and diseases by region and by bivalve type (nd indicates that pa­ rameter was not detected in a given bivalve species) based on the 2008 - 2009 MWP data.

Parameter American American Blue mussel Blue mussel Zebra mussel

oyster (Gulf) oyster (East) (East) (West) (GL) Synthesis Prokaryotes 1.09 2.36 0.60 0.39 nd Gregarines 38.20 20.12 0.08 4.65 nd H. nelsoni nd 2.18 - - - P. marinus 1.59 1.34 - - - Ciliates 1.87 8.76 3.08 2.23 nd Cestodes 0.56 0.72 nd nd nd Trematodes 0.15 0.06 2.88 0.07 nd Nematodes 2.12 1.84 nd nd 1.00 Copepods 0.06 0.04 0.12 0.41 0.00 Pea crab 1.00 1.00 0.93 0.0 - Hemocytic infil­ 0.45 1.00 1.73 0.69 0.19 tration Ceroid body 62.03 74.52 15.35 9.35 8.22 Digestive gland 2.10 2.38 2.54 2.63 1.86 atrophy Necrosis 0.00 0.00 0.41 0.07 0.02 Xenoma 2.79 2.00 2.77 nd

1992). These dreissenid mussels were introduced from parasite infection and disease follow the trend, from Europe (Hebert et al., 1989) and quickly invaded the highest to lowest: Gulf coast oysters > East Coast oys­ Great Lakes system, spreading through the lakes and riv­ ters > East Coast blue mussels > West Coast blue mus­ ers, except Lake Superior (Griffiths et al., 1991 and Mills sels > Great Lakes zebra mussels (Tables 4 and 5). et al., 1996). The lower prevalence of parasitism and dis­ ease in zebra mussels (Great Lakes) relative to the other The susceptibility of bivalves to harbor parasites bivalve species may likely be a corollary of the invasive was assessed using parasite taxa richness. Parasite taxa nature of the zebra mussels because ciliates, at least, are richness was calculated as the total number of parasite well known parasites in these species’ native waters (Kara­ types found in a bivalve species. Parasite taxa richness tayev et al. 2002). However, individual parasitic taxa that for 2008 - 2009 differed broadly among host bivalves did infect the dreissenids were also low in prevalence and and between geographic locations (Figure 62). Annual in infection intensity, suggesting that dreissenid popula­ parasite taxa richness in oysters varied from 2.5 in Ha­ tion dynamics might also be important. High growth rates waii to 17.9 in the Gulf of Mexico (Appendix B). With and short life spans offer obvious mechanisms to mini­ an annual average of 12.2, West Coast blue mussels had mize prevalence and infection intensity of parasites. The significantly higher (p < 0.05) parasite taxa richness than newness of introduced species in an ecosystem is often East Coast mussels, which had an annual average of linked to low parasite infection (Torchin et al., 2002 and about 9.8 (Appendix B). The American oyster harbored Marshall et al., 2003). significantly more parasite taxa than the other bivalves studied (p < 0.0001). Gulf coast oysters had significant­ Lower prevalence values for the histopathology param­ ly more diverse parasite types than East Coast oysters, eters in zebra mussels, therefore, should not be construed which in turn harbored more parasite types than oysters to imply that the Great Lakes have better water quality from Hawaii and Puerto Rico (p < 0.0001). Among all than the nation’s coastal and estuarine areas. In general, sentinel bivalve types, zebra mussels had the least di­

41 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

Table 6. Comparison of prevalence of histopathology parameters among bivalve species from 2008 - 2009 using analysis of variance (ANOVA). For each parameter, bivalves that do not have the same letter are statistically different at p < 0.05 (nd = parameter was not detected).

Synthesis Parameter American Blue mussels Blue mussels Zebra oysters (west) (east) mussels Parasite Prokaryotes A B A nd Gregarines A B C nd H. nelsoni A - - - P. mar inu s A - - - Ciliates B C A nd Cestodes A nd nd nd Trematodes B B A nd Nematodes A nd nd B Copepods A A A A Pea crab A B AB - Disease Hemocytic B BC A C infiltration Ceroid body A B BC C Digestive gland B A A B atrophy Necrosis B AB A B Xenoma A AB B nd

verse parasite types, with an annual average of 1.1 (p < stress response in zebra mussels and blue mussels, and 0.0001). stress could increase the susceptibility to opportunistic parasite infection. However, in this study, significant cor­ Over the monitoring period (1995 – 2009), annual average relations between contaminants and histopathology pa­ parasite taxa richness in Gulf Coast oysters was fairly con­ rameters were rare (Appendix A). Several reports based stant, except for the years between 2003 and 2006, where on the NS&T monitoring data (Apeti et al., 2009 and the values fluctuated (Figure 63a). The most variation in 2012; Kimbrough et al., 2008) indicated that heavy met­ parasite taxa richness was recorded in the blue mussels al and organic contaminant body burdens in the sentinel (Figure 63b). Annual averages of parasite taxa richness in bivalves were often very low (relative to U.S. Food and northeast bluemussels varied between 6 and 13, while those Drug Administration guidelines), which may explain the in West Coast blue mussels fluctuated between 10 and 16. lack of strong correlations with the histopathology pa­ Except for a few instances (1998, 2007 and 2008) where rameters in this study. Thus, the variation in parasite taxa there were no overlaps, parasite taxa richness appeared richness among bivalve species, as well as the presence to fluctuate similarly in blue mussels from both coasts. of parasite infection and disease hot-spots observed at various monitoring sites, may be attributable to an amal­ Exposure to contaminants is known to impact aquat­ gam of stressors, including biological and physical factors. ic bivalve immune systems and facilitate parasitism and The NS&T program assays different mollusks with dis­ prevalence of disease (Weis et al., 1995; Johnson et similar habitats, feeding rates, and physiological processes, al., 1992; MacKenzie et al., 1995). Rocher et al. (2006) all of which could contribute to their predisposition to showed that contaminants, including metals, activate disease and parasite infection. Aside from host biological 42 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters Table 7. Comparison of prevalence of histopathology parameters in bivalve among coastal regions from 2008 - 2009 using analysis of variance (ANOVA). For each parameter, coastal regions that do not have the same letter are statistically different at p < 0.05 (nd = parameter was not detected).

Parameter Southeast GOM Northeast Mid-Atlantic West coast Great

Lakes Synthesis Parasite Prokaryotes A BC BC AB C nd Gregarines A A C BC B nd P. marinus A A - B - - H. nelsoni A nd - A - - Ciliates AB BC A B BC nd Cestodes A B nd B nd nd Nematodes B A nd B nd Trematodes C C A B C nd Nematodes B A nd B nd Copepods A A A A A A Pea crab A A A A A - Disease Hemocytic ABC AB A A BC C infiltration Ceroid body A A B B B C Digestive BC C AB A A C gland atrophy Hemocytic ABC AB A A BC C infiltration Necrosis AB B AB A AB AB Xenoma B A B B B nd factors, environmental factors, such as water quality, sea­ disease emergence in marine and coastal environments has sonal fluctuation of temprature, and runoff, may affect been linked to climate change, which is shifting the dis­ a bivalves’ susceptibility to parasitism and disease. Water ease landscape globally (Harvell et al., 2002). The NS&T quality parameters, including coastal flow regime, water MWP collects samples only during the winter months residence time, and, more importantly, surface tempera­ (Apeti et al., 2011), hence, the dataset is not conducive tures and salinity, are known to have impacts on parasite to assess seasonal fluctuations of parasitism and disease. infection in bivalves (Ward and Lafferty, 2004). MacKen­ Outbreaks of disease and parasites could have potential zie et al. (1995) discussed how stressors, such as increased human health and economic consequences. Bivalves, such surface temperature and low dissolved oxygen, impact the as American oysters, are important commercial fisheries; physiological processes of bivalves and increase their sus­ outbreaks of pathogens such as P. marinus and H. nelsoni, ceptibility to infection. Temperatures above thermal tol­ can reduce stocks as a result of massive die-offs (Ewart erance were also shown to significantly stress organisms, and Ford, 1993). Other parasites, such as PIB (chlamydia increasing their susceptibility to some diseases, especially and rickettsia), cestodes (tapeworms), nematodes (round­ in aquatic ecosystems (Lafferty et al., 2004; Robinson et worms) and trematodes (flukes), only parasitize mollusks al., 2005). Thus, the increasing threat of coastal condition as juveniles, while their adult forms often live in the di­ changes resulting from climate change constitute poten­ gestive tract of vertebrates, like humans, where they can tial stressors for marine and coastal resources (Cook et al., cause serious health issues (Bogitsh and Carter, 2005). 1998). Recently, a number of biochemical alterations and

43

NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program Synthesis

25

20

15

10 P ar as i t i c t ax a r i c hnes s hnes c i r a ax t c i t i as ar P

5

0

oyster PR oyster HI oyster East oyster Gulf zebra mussel mussel East mussel West

Figure 62. Parasite taxa richness in different host bivalve types by region. The upper and bottom limits of the boxplots represent the 25th and 75th percentiles, the whiskers represent the 5th and 95th percentiles, and the line in the middle of the box is the median.

44

Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters Synthesis

Figure 63a. Temporal variation of parasite taxa richness in oysters from the East Coast and the Gulf of Mexico from 1995 - 2009.

18 16 14 12 10 8 6

Parasite taxa richness taxa Parasite 4 2 0

mussel East coast mussel West coast

Figure 63b. Temporal variation of parasite taxa richness in blue mussels from the East and West Coasts of the U.S from 1995 - 2009.

45 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program CONCLUSION

The status and trends of parasites and disease in sentinel bivalves (oysters and mussels) monitored by the NOAA/ NCCOS Mussel Watch program were assessed. Results indicated that parasites and disease were widespread, but Conclusion at variable prevalence and intensity. Parasites and disease were found to be generally low, although some hot-spots of high pevalence values were observed at locations around the U.S., especially at oyster sites. Overall, prev­ alence and intensity of parasites and disease showed the following regional trend, from highest to lowest: Gulf Coast oysters > East Coast oysters > East Coast blue mussels > West Coast blue mussels > Great Lakes zebra mussels. Zebra mussels harbored fewer parasites, perhaps due to the fact that they were recently introduced to the Great Lakes and as a result are not being used as interme­ diate hosts. It should be noted that there are other histo­ pathology parameters in shellfish that the MWP does not assess. These include, amongst others, paralytic shellfish poisoning and fecal coliform bacteria, which both pose significant threats to human health.

Parasite infections and disease have been linked to con­ taminants in water, but this study showed little relationship between parasites and disease in bivalves and their body burden of chemical contaminants. Additionally, temporal trends assessment indicated that parasite infections and presence of disease were fairly static over the monitoring period assessed in this study, although in a few instances some site-specific decreasing and increasing trends were observed. Both the temporal trends and correlation be­ tween histopathology parameters and contaminants were not extensive enough to infer conclusion on water quality.

Environmental stressors, including anthropogenic events, climate change, and the resulting impacts of sea level rise and increased surface temperature, will continu­ ously influence the distribution of disease and pathogens in our coastal waters. Because multiple species belonging to various coastal zones were used, in case of unforeseen events, such as outbreaks of shellfish disease or pathogens, the NS&T MWP would be unique in providing baseline histopathology data to coastal resource managers.

46

Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters Conclusion

47 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

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51 NATIONAL STATUS AND TRENDS PROGRAM: The Mussel Watch Monitoring Program

Appendix A Correlation between parasitic taxa and contaminants measured by the NS&T Mussel Watch Pro­ gram. Results represent pair of parameters, which displayed statically significant relationships (p <

0.05). Appendices

Bivalve Histopathology Contaminant Spearman ρ P value species parameter American oyster Cestodes Arsenic 0.492 < 0.001 American oyster Cestodes Cadmium -0.422 < 0.001 American oyster Cestodes Nickel -0.377 < 0.001 American oyster Nematodes Cadmium 0.300 < 0.001

Blue mussels Gregarines Arsenic 0.329 < 0.001 (West) Blue mussels Gregarines Lead 0.315 < 0.001 (West) Blue mussels Gregarines Manganese -0.343 < 0.001 (West) Blue mussels Ciliates Manganese 0.349 < 0.001 (West)

Blue mussels Ceroid body Arsenic 0.369 < 0.001 (West) American oyster Gregarines Cestodes 0.306 < 0.001 American oyster P. marinus Gregarines 0.307 < 0.001 Blue mussels Digestive gland Necrosis 0.338 < 0.001 (East) atrophy

52 Parasites and Disease in Oysters and Mussels of the U.S. Coastal Waters

Appendix B Annual parasite taxon richness by bivalve type and geographic location.

Oysters Zebra Blue Mussels Appendices Mussels Year East Gulf Puerto Hawaii Great East West Alaska Rico Lakes 1995 16 17 1 11 16 7 1996 16 19 6 9 12 1997 18 19 2 9 14 3 1998 15 18 9 1 1 12 12 1999 17 18 7 13 3 2000 14 19 7 1 8 14 2001 17 18 1 6 10 3 2002 16 18 5 4 9 10 2003 15 16 12 11 4 2004 13 20 3 1 10 14 2005 15 15 1 9 12 5 2006 14 19 6 2 11 11 2007 14 17 13 11 4 2008 14 17 6 9 12 3 2009 17 18 1 12 11 3 Average 15.4 17.9 6.5 2.5 1.1 9.8 12.2 3.9

Data Access Unlike the chemistry data that is publically available through the NS&T data portal (http://egisws02.nos.noaa.gov/ nsandt/index.html#), the NS&T histopathology data is only available by request. Please call (301) 713-3028.

53

U.S. Department of Commerce Penny Pritzker, Secretary

National Oceanic and Atmospheric Administration Kathryn Sullivan, Under Secretary for Oceans and Atmosphere

National Centers for Coastal Ocean Science: Delivering ecosystem science solutions for stewardship of the nation’s ocean and coastal resources, in direct support of National Ocean Sevice priorities, offices and customers to sustain thriving coastal communities and economies.For more information, visit: http://www.coastalscience.noaa.gov/.