Canadiantechnical Report of Fisheries and Aquatic Sciences No

59 Shell Disease (Ostracoblabe implexa)

Background Information - This disease ("Dutch Shell disease", "Maladie du pied" or "Maladie de la charnière") affects edible oysters (Ostrea edulls) wherever grown in Atlantic Canada, as well as in Europe. It was introduced to Atlantic Canada in 1969 with broodstock from Connecticut which were held in quarantine for breeding. Subsequent culture of edible oysters in the Gulf of St. Lawrence in the 1970's was thwarted by proliferation of the fungal agent of shell disease (Ostracoblabe implexa) during the warm summer water conditions. Along the Atlantic coast of Nova Scotia high temperatures do not last long enough for the disease to become a significant problem. No other bivalves are affected, even when grown be,side heavily infected O. edulis stocks.

Effect on Host - Ostracoblabe implexa proliferates through the shell and may penetrate to the inner surface where white spots and conchiolin warts are formed. Shell closure is impeded when the fungus affects adductor muscle attachment or causes thickening of the shell margin and hinge ("maladie de charnière"). In severe infections the oyster is weakened and dies.

Commercial Significance - The effects of this disease have made edible oyster culture in PEI waters uneconomical. Although no mass mortalities have been attributed to shell disease along Scotian shelf waters, shell deformities can reduce the market value of the oyster.

Diagnosis - Gross diagnosis: Shell deformities, as described above. (Figure 27A). Histology: Presence of aseptate mycelia in decalcified shell sections. Squash preparations: Observation of dense mycelial networks of straight hyphae (1.5- 2.5 iLm in diameter) with oval dilations at irregular intervals (40-100 gm) (Figure 27B). Septa may be seen on dying mycelia. Culture: Shell fragments, incubated at 25°C for —5 weeks in sterile seawater with antibiotic, develop mycelial threads that can be cultured on yeast peptone agar at 25°C.

Prevention - Stocks with a history of shell disease should not be introduced into areas where water temperatures exceed 22°C for more than two weeks. Any calcareous-based substrate will act as a reservoir for the fimgus, which makes subsequent eradication difficult.

References - Alderman, D.J. 1986. Fungal disease,s of marine invertebrates. European Aquaculture Society, Special Publications 9:197-201.

Alderman, D.J. and E.G.B. Jones. 1971. Physiological requirements of two marine phycomycetes, Althornia crouchii and Ostracoblabe implexa. Transactions of the British Mycological Society 57:213-225.

Li, M.F., R.E. Drinnan, M. Drebot and G. Newkirk. 1983. Studies on shell disease of the European flat oyster Ostrea edulis Linné in Nova Scotia. Journal of Shellfish Research 3:135-140. (continued on page 61)

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Figure 27: (A) Shell damage to an edible oyster (O. edulis), infected by the fungus Ostracoblabe implexa. Note the extensive dark discolouration of the calcareous layer underlying the thin nacre layer of the shell and extensive shell "wart" formation (arrows) around the adductor attachment area of the shell. The hinge and edge of the shell also show characteristic thickening. (Material provided by Dr. D. J. Alderman). (B) Hyphal network of Ostracoblabe implexa cultured from infected edible oyster (O. edulis) shell. (Scale bar = 50 iim). (Material provided by Mr. R.E. Drinnan). 61

Hexamitiasis

Background Information - Hexamita inflata (Protozoa: Sarcomastigophora) is a flagellate protozoan commonly found in tissues of dying e,astern oysters (Crassostrea virginica) as well as healthy overwintering oysters throughout Atlantic Canada and the eastern United States. The saine (or a similar) flagellate has also been found in moribund quahaugs (Mercenaria mercenaria) and edible oysters (Ostrea edulis). Oysters dying from Malpeque disease were found to be infected by Hexamita, however, the same flagellate was isolated from the blood of neighbouring, healthy, oysters. Unlike some other Hexamita species (e.g. H. salmonis of salmon), H. inflata is a saprobiont (organism which thrives on decaying tissue) rather than a true parasite (causing damage to healthy tissue). The Hexamita species which invade oysters are found free-living in anaerobic (oxygen depleted), muddy substrates. Hexamita species have also been reported in association with mortalities of oyster species elsewhere (Ostreola conchaphila (Olympia oyster) from Puget Sound, Washington, USA and O. edulis from the Netherlands) but their actual role in these mortalities was not determined.

Effect on Host - The direct effect of Hexamita on oysters is not clearly established. Although found at low intensities in moribund oysters, in association with necrotic tissue, studies have shown that this flagellate is not the cause of morbidity. The same flagellate is regularly found in the blood of healthy oysters in anaerobic winter conditions. Canadian studies suggest that this organism is an early saprobiotic invader of the tissues of oysters weakened or dying from other causes.

Commercial Significance - None known.

Diagnosis - Wet mount: Preliminary diagnosis can be made by examining a drop of oyster haemolymph for highly motile, pear-shaped, flagellates measuring 8-16 gm. (Figure 28A). Histology: Examination of tissue cross-sections for flagellated protozoans in tissue sinuse,s (Figure 28B). Identification to species re,quires stained smears of the flagellates (see Wet Mount above). Culture: Anaerobic culture using HM medium supplemented with beef serum and agar omitted (see Khouw, McCurdy and Drinnan (1968)).

Prevention - There is no known treatment. Prevention may not be practical due to the wide distribution of Hexaniita in muddy substrates and no clear evidence of direct pathology. 62 References - Feng, S.Y. and L.A. Stauber. 1968. Experimental hexamitiasis in the oyster Crassostrea virginica. Journal of Invertebrate Pathology 10, 94-110.

Khouw, B.T. and H.D. McCurdy. 1968. Nutritional studies of a flagellated protozoan Hexamita inflata from the Canadian oyster, Crassostreavirginica. Canadian Journal of Microbiology 14:817-821.

Khouw, B.T., McCurdy, H.D.,Jr. and R.E. Drimian. 1968. The axenic cultivation of Hexamita inflata from Crassostrea virginica. Canadian Journal of Microbiology 14:184- 185.

Scheltema, R.S. 1962. The relationship between the flagellate protozoan Hexamita and the oyster Crassostrea virginica. Journal of Parasitology 48:137-141.

Schlicht, F.G. and J.G. Mackin. 1968. Hexamita nelsoni sp.n. (Polymastigina: Hexamitidae) parasitic in oysters. Journal of Invertebrate Pathology 11:35-39.

Stein, J.E., J.G. Denison and J.G. Mackin. 1959. Hexamita sp. and an infectious disease in the commercial oyster Ostrea lurida. Proc. of the National Shellfisheries Association 50, 67-81.

(continued on page 64) 63

Figure 28: (A) Hexamita sp.collected from a culture of necrotic quahaug tissues (M. mercenaria) (Scale bar = 100 gm). (B) Necrotic mantle tissue of an eastern oyster (C. virginica) showing aggregations of Hexamita sp. (arrows) in the connective tissue and in the haemolymph, necrotic haemocytes, as well as connective and epithelial tissue breakdown (H&E, Scale bar = 20 gm). 64

MUSSEL PARASITES, PESTS AND DISEASES

Haemic Neoplasia

Background Information - Neoplasias have been reported from fifteen bivalve species around the world. The most common type of neoplasia affects the blood cells ("haemic neoplasia"), as is the case for blue mussels (Mytilus eclair's) from North America and Europe. In Atlantic Canada low prevalences of this disease have been observed, and none have been associated with mass mortalities, as reported for a similar neoplasia in soft-shell clams (Mya arenatia) from Chesapeake Bay, USA. Scientists from the west coast of the United States have transmitted haemic neoplasia from infected to uninfected blue (bay) mussels under laboratory conditions. Similar transmission studies with mussels from Atlantic Canada failed. The cause of haemic neoplasias has been related to a number of different factors (viral infections, pollution effects or genetic susceptibility). Both suspension- and bottom- grown populations are affected with prevalence,s peaking in late summer and early autumn.

Effect on Host - Pathogenicity is linked to disruption of normal cell-function. The nuclei of affected blood cells are diffuse and enlarged (compared to their normal condensed and granular appearance), numbers of blood cells increase, as does cell division (mitosis). Differences in levels of mortality between stocks may indicate acquired disease resistance, a genetic component to susceptibility or multiple pathogenicity factors.

Commercial Significance - Sporadic mussel mortalities around the Magdalene Islands and in mussels ftom Prince Edward Island, have shown high prevalences of haemic neoplasia. Similar neoplasia has also been associated with significant losses of blue (bay) mussel stocks on the Pacific c,oast. The cause/effect relationship, however, has not been clearly established.

Diagnosis - Histology - Presence of haemocytes with large diffuse nuclei and relatively little cytoplasm, compared to normal haemocytes. Advanced haemic neoplasia is characterised by extensive infiltration throughout the connective tissue, diapedesis and the presence of mitotic figures (compare Figure 29 with 30A, B). Early stages of the disease are less well defined and may appear as pockets of abnormal looking haemocytes with or without evidence of mitosis or undifferentiated nuclei (diffuse enlargement). (The germinal epithelial cells of the gonoducts normally show features characteristic of neoplastic cells, i.e., mitotic figures and undifferentiated nuclei). Blood Smear - Presence of blood cells showing the features noted above for histology. Severely affected haemocytes lack the capability to adhere to the microscope slide and tend to round up. I I 65 Prevention - It is advisable to avoid introduction of stocks demonstrating haemic neoplasia into areas I where neoplasia has not been detected previously or is not considered a significant disease problem.

I References - Bower, S.M. 1989. The summer mortality syndrome and haemocytic neoplasia in blue mussels (Mytilus edulis) from British Columbia. Canadian Technical Report of Fisheries I and Aquatic Sciences 1703:1-65. Elston, R.A., M.L. Kent and A.S. Drum. 1988. Progression, lethality and remission of I hemic neoplasia in the bay mussel, Mytilus edulis. Diseases of Aquatic Organisms 4: 135-142. I Farley, C.A. 1976. Proliferative disorders in bivalve mollusks. Marine Fisheries Review 38(10): 30-33. I McGladdery, S.E. and M.F. Stephenson. 1991. Parasites and diseases of suspension- and bottom-grown shellfish from eastern Canada. Aquaculture Association of Canada Bulletin I 91-3: 64-66. Peters, E.C. 1988. Recent investigations on the disseminated sarcomas of marine bivalve I molluscs. American Fisheries Society Special Publication 18: 74-92. I I I I t I I I (continued on page 68) I I 66

Figure 29: Haematoxylin and eosin stained tissue-section through the tissues of a healthy blue mussel (Mytilus edulis). Mantle (m), gonad (gd), gills (gl), digestive gland (dg), stomach (s), intestine (i), foot-retractor muscles (rm) and foot (f) (Scale bar = 2 mm).

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haemocytes (n) are present among the neoplastic cells. (dt - digestive tubule, int - intestine) (H&E, Scale bar = 100 um). I 68

Gill Ciliates of Mussels (Ancistrum mytili)

Background Information - I Ancfstrum mytili (Protozoa: Ciliata: Kinetofragminophora) ciliates have been reported on the gills of blue mussels (Mytilus edulis) from around the world. They are commonly found on both suspension- and bottom-grown mussels from Atlantic Canada with numbers I of over a hundred occurring in a single tissue-section. A. mytili vary in size and shape and may or may not be attached to the gill surface. Additionally, some of these gill ciliates I may be infected by a rickettsial-like "hyperparasite" (parasite of a parasite). Effect on Host - There is no obvious host response to A. mytili, although some scientists have speculated I that large numbers may impair normal feeding and respiration under adverse growing conditions. There has been no evidence of this, to date, in Canadian Atlantic mussels. I Commercial Significance - None known. I Diagnosis - Gross Observation: Presence of ciliates (barely visible to the naked eye) on the surface of the gills and mantle. I Gill Smear: Ciliates collected from gill smears should be stained using a silver stain to check the species identification, where necessary. Histology: Ciliates on and between the gill lamellae, characterised by a large kidney- I shaped granular macronucleus, a smaller less basophilic micronucleus, numerous vacuoles, a pyriform body shape and a non-uniform covering of cilia (Figure 31A). Body size ranges from 30 µm to 60 µm. Rickettsial-like hyperparasites (Figure 31B) may be I present and appear as small, dark staining (blue-black), inclusions inside the ciliate. Prevention - These ciliates have not been clearly associated with pathology. Higher numbers occur on 1 bottom-grown mussels, compared to suspension-grown mussels, so mussel lines stocked with spat collected from suspension (rather than from bottom-grown stock) may help I prevent accumulation of potentially harmful ciliate numbers. References - I Fenchel, T. 1965. Ciliates from Scandinavian molluscs. Ophelia 1:113-120. Hatzidimitriou, G. and J. Berger. 1977. Morphology and morphogenesis of Ancistrum mytili (Scuticociliatida: Thigmotrichina), a commensal ciliate of mytilid pelecypods. I Protistologica 13: 477-495. McGladdery, S.E. 1990. Shellfish parasites and diseases on the east coast of Canada. I Aquaculture Association of Canada Bulletin 90-3: 14-18. McGladdery, S.E. and M.F. Stephenson. 1991. Parasites and diseases of suspension- and I bottom-grown shellfish from eastern Canada. Aquaculture Association of Canada Bulletin 91-3: 64-66. I (continued on page 70) I 69

Figure 31: (A) Ancistrum nwtili ciliates (arrow) on the gills of a blue mussel (M. edulis). Note the loose association between the ciliates and the gill epithelium (compare with the Sphenopluya-like ciliates of eastern oyster (Figure 22B) and blue mussel (Figure 32B). (H&E, Scale bar = 100 gm). (B) Specimen of A. mytili containing rickettsial-like hyperparasites (arrow) (H&E, Scale bar = 20 gm). I 70 P Gill Ciliates of Mussels (Sphenophrya-like)

Background Information - I Sphenophrya-like ciliates (Protozoa: Ciliata: Kinetofragminophora) are found attached to the gills of bottom- and suspension-grown mussels (Mytilus edulis) throughout Atlantic Canada. They usually occur in low numbers (1-2 per tissue-section) and are not I considered to be pathogenic. Similar ciliates have been described from Cmssostrea virginica (page 49) and have occasionally been reported as causing massive enlargement of infected gill cells and their nuclei ("xenoma" - parasite-induced hypertrophy) in eastern I US populations. This type of gill lesion has been found once on blue mussels from Atlantic Canada, but not yet on C. virginica. I Effect on Host - Usually there is no obvious host response to these ciliates, even when embedded in the gill or palp epithelium. A single specimen with ciliate-induced xenomas (host cell I hypertrophy) has been observed in a healthy mussel from the Gulf of St. Lawrence. Whether or not this xenoma was caused by the same species of ciliate found on the gill I surface, however, was not determined. Commercial Significance - I None known. Diagnosis - Histology: Presence of small, pear-shaped, ciliates (18 µm to 24 µm long) attached to I gill or palp epithelia (Figure 32A). The anterior (pointed) end is usually attached to the cell surface. Xenoma formation (Fig 32B) is rarely observed.

Prevention - I Since these ciliates generally occur in low numbers with no distinct host-response, prevention appears unnecessary.

I References - Fenchel, T. 1965. Ciliates from scandinavian molluscs. Ophelia 2(1): 71-174 I (descriptions of Sphenophrya spp. from Scandinavian clam species only). McGladdery, S.E. and M.F. Stephenson. 1991. Parasites and diseases of suspension- and bottom-grown shellfish from eastern Canada. Aquaculture Association of Canada Bulletin 91-3: 64-66.

Otto, S.V., J.C. Harshbarger and S.C. Chang. 1979. Status of selected unicellular I eucaryote pathogens, and prevalence and histopathology of inclusions containing obligate prokaryote parasites in commercial bivalve mollusks from Maryland estuaries. Haliotis 8: I 285-295. Sprague, V. 1970. Some protozoan parasites and hyperparasites in marine bivalve molluscs. In: Snieszko, S.F. (ed) A Symposium on Diseases of Fishes and Shellfishes. I American Fisheries Society Special Publication No. 5: 511-526. I (continued on page 72) I 7

Figure 32: (A) Sphenopiuya-like ciliate attached to the gills of a blue mussel (M edulis). Compare with Sphenopluya-like ciliates of eastern oyster (Figure 22B). (H&E, Scale bar = 100 gm). (B) Sphenopluya-like ciliates inside an enlarged gill epithelial cell. Note distension of the cell nucleus (hn) as well as the cell itself. The ciliates can be seen with tentacle-like (arrows) filamentous extrusions into the host cell cytoplasm. (H&E, Scale bar = 100 ..tm). 72

Digestive Tubule Ciliates

Background Information - "MPX" ("mussel protozoan X", after Bower, S. M., pers. comm.) is an intracellular ciliate which occurs in the digestive tubules of blue mussels (Mytilus edulis) from North America and Europe. They are found buried inside the epithelial cells or, more rarely, partially embedded or outside the host epithelium. In heavy infections (>500 per tissue- section) more than one ciliate may occur inside a single cell, however, normal levels range from 5-50 per tissue-section. The exact identity of the ciliate is still being studied but is believed to belong to the Class Kinetofragminophorea (shnilar to the ciliates described from the gills and tubules of mussels and other Atlantic bivalves).

Effect on Host - Although MPX causes cell damage, no specific host response has been observed, even in tissue-sections with over 1000 MPX.

Commercial Significance - None known.

Diagnosis - Histology: Observation of protoz,oans lying within or partially embedded in the digestive tubule epithelial cells (Figure 33A). Although small and intracellular (MPX ranges in length from 6 gm to 12 gm, compared to the host cell which varies from 30 tan to 40 gIll in height), they are easily detected by their dense macronuclei which are polymorphic (most commonly globular) and stain deep blue (densely basophilic). The tubule epithelial cells have less dense, granular, nuclei. The cilia of these ciliates are not easily seen except under high magnification (oil immersion). MPX maintain their cilia within the host cell (Figure 33B) (unlike the Sphenophrya-like ciliates described on page 70) and most are surrounded by a translucent "vacuole" within the host cell.

Prevention - Due to the lack of pathology associated with this ciliate, and its ubiquitous distribution throughout the range of the blue mussel, prevention appears unnecessary.

References - Figueras, A.J., C.F. Jardon and J.R. Caldas, 1991. Diseases and parasites of mussels, Mytilus edulis, Linnaeus 1758) from two sites on the east coast of the United States. Journal of Shellfish Research 10: 89-94.

McGladdery, S.E. 1990. Shellfish parasites and diseases on the east coast of Canada. Aquaculture Association of Canada Bulletin 90-3: 14-18.

McGladdery, S.E. and M.F. Stephenson. 1991. Parasites and diseases of suspension- and bottom-grown shellfish from eastern Canada. Aquaculture Association of Canada Bulletin 91-3: 64-66.

(continued on page 74) 73

Figure 33: (A) "MPX" (mussel protozoan X) ciliates (arrows) inside the digestive tubule (dt) cells of a blue mussel (M. edulis). (H&E, Scale bar = 0.5 mm). (B) "MPX" ciliates lying in and between the tubule epithelial cells. Note dark staining, macronucleus (pn) compared to the granular nucleus of the tubule cells (en). Note also the vacuole-like space (vs) around many of the ciliates inside epithelial cells. (H&E, Scale bar = 20 gm). I I 74 Intestinal Ciliates of Mussels

Background Information - I Ciliate protozoans are found in the intestine of both bottom- and suspension-grown blue mussels (Mytilus edulis) from Atlantic Canada. They are similar in appearance to Ancistrwn mytili (see Gill Ciliates of Mussels, page 68) but less common (generally less I than 10% prevalence and less than 5 per tissue-section). This low prevalence means that neither the identity of the ciliate or any seasonal variation has yet been determined. They I lie between the food contents and wall of the intestine but do not appear to be attached. Effect on Host - To date, no infections have been associated with pathology and no host response has been I observed. Commercial Significance - I None Known. Diagnosis - I Histology: Observation of ciliates lying within the intestine, usually up against the epithelium (Figure 34A). They vary in size from 36 µm to 42 µm in length, although measurements may be significantly altered in specimens collected whole. They are I characterised by granular cytoplasm and densely basophilic (blue-staining) macronucleus (Figure 34B). Their body shape moulds to the shape of the intestine wall, as well as to any neighbouring ciliates, giving a mosaic-like appearance when more than one is present. They appear to have a uniform distribution of cilia but differ from A. mytili in shape and A the lack of distinct food vacuoles. There is no evidence of a suctorial "tentacle" or hyperparasitic rickettsial-like inclusions.

I Prevention - Low prevalences and lack of specific identification for this ciliate makes prevention impractical, to date. Furthermore, prevention may be unnecessary, since there is no I evidence of associated pathology.

References - I McGladdery, S.E. and M.F. Stephenson (Unpublished data). I I I

I (continued on page 76) I I 75

Figure 34: (A) Ciliates (arrows) inside the intestine of a blue mussel (M edulis) (H&E, Scale bar = 100 gm). (B) Intestinal ciliates showing large basophilic nuclei and polymorphic body shapes. Note the lack of attachment to the intestinal epithelium demonstrated by other internal ciliates (Figures 23A, B, and 33A, B). (H&E, Scale bar = 20 gm). I 76 I Digenean metacercariae Background Information - Digenean larvae (metacercariae) are commonly found in bottom-growing mussels (Mytilus I edulis) throughout Atlantic Canada. They encyst in the mantle, foot and digestive gland where the host may surround them with a pearl. Infections are usually low (2-5 per tissue- section) in bottom-growing mussels and rare in suspension-grown mussels. Although I difficult to identify from tissue sections, small anterior spines are sometimes visible indicating that they are Echinostomatid or Gymnophallid digeneans. I Effect on Host - Mussels control tissue damage by digenean metacercariae by "walling them off" inside pearls. Despite tissue disruption directly around the metacercariae, neighbouring tissues I appear unaffected. Commercial Significance - I The presence of pearls in the soft-tissues of mussels may reduce marketability. Consumers prefer to wear pearls rather than eat them. Since the pearls encountered in Atlantic blue mussels are usually small (less than 1-2 mm long), vary from white to grey and are rarely I spherical, they have no commercial value. Diagnosis - I Gross Observation: Observation of pearls of various sizes on the mantle indicates the possible presence of digenean metacercariae (Figure 16C). Note -pearls may be initiated by any irritant too large to be expelled by host haemocytes. Histology: Metacercariae occur in the connective tissue of the mantle, gills and digestive I gland where they are usually surrounded by intense haemocyte infiltration and an eosinophilic (pink-staining) capsule (the pearl) (Figure 35A). Some, however, may be found with no granuloma (Figure 35B). Metacercariae can be differentiated from other r foreign objects stimulating pearl-formation by the presence of body-parts (including suckers, spiny integument, etc) or a distinct vacuole in the centre of the pearl. Note - Pearls may be impede sectioning of tissue-blocks. Chipping of microtome blades and I consistent tears along the ribbon indicate calcareous inclusions in the tissues. Heavily infected tissues can be fixed in an acidic fixative (e.g. Davidson's) and/or declacified in I EDTA (ethylenediaminetetraacetate) post-fixation and prior to paraffin embedding. Prevention - Metacercarial infections can be reduced by suspension-culture away from the infective I cercarial stages released from benthic gastropods, especially during peak cercarial release over the summer months. (Caution should also be taken to reduce mussel exposure to I other tissue irritants). References - Lauckner, G. 1983. Diseases of mollusca: Bivalvia. pp. 632-762. In: Kinne, O. (ed) I Diseases of Marine Animals, Vol II. Biologische Anstalt Helgoland, Hamburg. McGladdery, S.E. and M.F. Stephenson. 1991. Parasites and diseases of suspension- and I bottom-grown shellfish from eastern Canada. Aquaculture Association of Canada Bulletin 91-3: 64-66. I (continued on page 78) I

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Figure 35: (A) Granuloma (g) associated with a pearl (p) encapsulating a digenean metacercaria (m) in the connective tissue of a blue mussel (M edulis). (H&E, Scale bar = 100 pin). (B) Pearl containing a digenean metacercaria in the connective tissue of a blue mussel (M edulis) but with no associated granuloma (H&E, Scale bar = 100 pm). I I 78 Tissue crustaceans I Background Information - Very little information is available on the presence of nauplii larvae and copepod-like crustaceans in blue mussel (Mytilus edulis) tissues. As with digenean metacercariae (page 76), these crustaceans can elicit extensive granuloma formation, however, they do not I appear to stimulate pearl formation. Older lesions containing necrotic crustacean debris develop into abscesses. The factors behind how or why these crustaceans end up in the mussel tissues has not been clearly determined, however, these types of lesions are found I almost exclusively in bottom-grown mussels throughout Atlantic Canada.

Effect on Host - I Crustacean tissue inclusions usually range from 1-2 per tissue-section, however, multiple lesions may take up over 50% of the mussel's connective tissue. Despite this intense I haemocyte response, no such infections have been associated with mussel mortality. Commercial Significance - t None known. Diagnosis - Histology: Observation of large granulomas or abscesses containing parts of crustaceans I (Figure 36A, B). Granulomas are commonly formed in mussels in response to other tissue irritants, so the presence of crustacean pleopods, antennae, etc., are necessary for verification of the causative agent. Note: Tissue-embedded crustaceans should not be I confused with either: the endoparasitic copepod of the mussel intestine, Mytilicola intestinalis ("Red-worm"), or "pea-crabs" (Pinnotheres spp) from the mantle cavity, I neither of which have been found to date in Canadian Atlantic waters. Prevention - Suspension-grown mussels are significantly less infected than bottom-grown mussels.

I References - McGladdery, S.E. and M.F. Stephenson. 1991. Parasites and diseases of suspension- and bottom-grown shellfish from eastern Canada. Aquaculture Association of Canada Bulletin I 91-3: 64-66. I I I

I (continued on page 80) I I

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Figure 36: (A) Copepod-like crustacean (c) in the connective tissue of a blue mussel (M edulis). Note the extensive granuloma (g) surrounding the crustacean, as well as throughout the digestive gland (arrows) (H&E, Scale bar = 0.5 mm). (B) Haemocyte response to a "stray" copepod within the tissues of a blue mussel (M edulis). Note the granulocyte infiltration throughout the necrotic crustacean debris. (H&E, Scale bar = 100 mn). 80

CLAM PARASITES, PESTS AND DISEASES

"QPX" disease

Background Information - In the early 1960's a mass mortality of quahaugs (hard-shell clams), Mercenaria mercenaria, from Neguac, New Brunswick, was attributed to an Olpidium-like chytrid fimgus ("Chytrid-disease" of Drinnan and Henderson, 1963). Subsequent mortalities occurred on PEI and more recently at a shellfish hatchery on PEI. To date, this organism has only been found in Canadian quahaugs. The causative organism has characteristics similar to Labyrhuhuloides haliotidis from abalone, Hanods spp. on the west coast of Canada (Bower, 1987), as well as certain perkinsiid apicomplexans (Whyte, S. and R. Cawthorn, pers. comm.), hence the use of the non-taxonomic name Quahaug Parasite "X" (QPX). Only certain stocks of adult quahaugs at the shellfish hatchery were killed, suggesting that some stocks may be more susceptible than others. The same organism has been found in healthy wild stocks, indicating that pathogenicity may also be linked to environmental factors.

Effect on Host - There is a massive haemocyte response to tissue invasion by QPX. Aggregations of "spore-like" stages are surrounded by necrotic haemocytes in the connective tissue of the digestive gland, mantle, gills, kidney and foot. Many of the parasites appear surrounded by a translucent "vacuole" indicating possible lysis of neighbouring host tissue. Mortalitie,s are believed to be due to proliferation of QPX throughout the tissues.

Commercial Significance - The 90-100% mortalities associated with QPX disease is a significant loss for both fishery exploitation and hatchery broodstock. No mass mortalities have been reported from the wild since the cases described from New Brunswick and PEI in the early 1960's, however, pathogenicity may be increased when infected quahaugs are kept in holding facilities.

Diagnosis - Histology: Presence of abscess lesions containing various stages of vegetative and spore- like spheres, commonly inside "vacuoles" of host tissue. An intracellular stage develops into an intercellular stage comprised of spheres (size range 12 km-60 gm). These spheres produce masses of daughter cells. Clear zones are visible around the spheres and daughter cells along with "stellar" (possibly cytoplasmic) extrusions from the vegetative stages. Some spheres are surrounded by a refractive cyst wall which ranges in width from 4-8gm (compare Figure 37 with Figure 38A, B). Tissue culture - QPX can be cultured from infected clam tissue samples on Potato Dextrose Agar. The cysts are retrieved from both the necrotic clam tissue (Figure 38C) and from fungal-like colonies which range in colour from cream or yellow. Motile spores may be observed in the cyst stage of QPX. I I 81 Prevention - Treatment and control have yet to be determined. Recent cases from Atlantic Canada I indicate it is highly infectious under certain conditions. Avoidance of importation of stocks carrying detectable levels of this organism, especially for hatchery use, is I advisable. References - Bower, S.M. 1987. Labyrinthuloides haliotidis n.sp. (Protozoa: Labyrinthomorpha), a I pathogenic parasite of small juvenile abalone in a British Columbia mariculture facility. Canadian Journal of Zoology 65:1996-2007. I Drinnan, R.E. and E.B. Henderson, 1963. 1962 mortalities and a possible disease organism in Neguac quahaugs. Annual Report Biological Station, St. Andrews, No.B11, I 3pp. McGladdery, S.E. and M.F. Stephenson, (unpublished data).

Whyte, S.K., R.J. Cawthorn and S.E. McGladdery. (in press) Pathology and in vitro development of an unidentified protistan parasite of the quahaug (Mercenaria mercenaria). I Diseases of Aquatic Organisms. I I A I I I I I i (continued on page 85) t 82

Figure 37: Haematoxylin and eosin stained tissue-section through the tissues of a healthy quahaug (hard-shell clam) (Mercenaria mercenaria). Mantle (m), gonad (gd), gills (gl), digestive gland (dg), stomach (s), intestine (i), kidney (k) and foot (f). (Scale bar = 0.2 mm). I

I 83 I I i I I I I ,,,* •. . • :,+,. -^ • .^. ;,. , w t !E * I I il J^+yir -il *.Il> - - . -0 • !T1 ,S' ^,R^ •s ^y (• il. ..`'.^ r '' i ^.:• ï^ ^^. o • ` ^ i _ • .;.. yfr,^I^ ^i 1

aaa.^^^^'N-&^M1???^ `^•. • i :^^• ^.,. â^^^._- • O.jM ^ , • ip b !^ `+ • ^ i âm* 1 ^; +.Yè^^ ^ t^ u^^+^, - ^ b^ ^ ^ •d ^ •+ • , I Ir • ., 0 • ^o ^" , •a*s 14i+1 ► ^ , . 2^^ r^ ^: • .• ^. ^ ^ :^1[•w^,^ ^•;^^ I ^.•s . •t^ ID

• ^ '• ^ ^ ^ ^ 7p.^^ ^ • : + ^ ^ ^^^• ^+ s ► I a^1^, ^ • r .^ i • . , .. ^. • ^ ^ ^iF ^^ ao • ^ .. d c^ • • • I li "P4 ^i $e i( " L` .-^ ^ •^ I --_ a o.* 0 Figure 38: (A) Cross-section through quahaug (M. mercenaria) showing lesions caused by a thraustochytrid-like organism "Quahaug Parasite X" ("QPX") (arrows). (m - mantle, dg - digestive gland, int - intestine). (H&E, Scale bar = 0.5 mm). (B) Multiplicative stages (ms) I and daughter cells (dc) of "QPX" in the mantle tissue of quahaug (M. mercenaria). (H&E, I Scale bar = 100 µm). I 84

Figure 38: (C) Colonies of Chytrid-like "Quahaug Parasite X" (QPX) cultured on Potato Dextrose Agar from necrotic quahaug (M mercenaria) tissue. Thick-walled, cyst-like stages (arrows) are interspersed among smaller "daughter cell" stages. (Scale bar — 100 gm). 85 Haemic Neoplasia

Background information - Haemic neoplasia of soft-shell clams (Mya arenaria) is also known as haematopoietic neoplasia and soft-shell clam sarcoma. It has been reported from Chesapeake Bay north to the Bay of Fundy, but has not yet been found in soft-shell clams from the Gulf of St. Lawrence. Mass mortalities have been attributed to this disease along the eastern US but no similar mortalities have been'observed in Canadian Atlantic populations. Prevalences in Atlantic Canadian clams appear to be low.

Effect on Host - Populations of soft-shell clams in Chesapeake Bay have suffered mortalities ranging from 30-80% since the early 1980s. Prevalence increases over winter with mortalities occuring during winter and early spring. Pathogenicity is due to disruption of normal cell-function. Soft-shell clams from the Bay of Fundy to Massachusetts have shown no evidence of mortality as a result of this disease. The variation in pathogenicity is still being studied.

Commercial Significance - This disease has no commercial significance to soft-shell clam populations in Atlantic Canada. The losses encountered in Chesapeake Bay, however, are highly significant.

Diagnosis - Histology - Detection of haemocytes with enlarged, diffuse, nuclei as well as evidence of mitosis and diapedesis (Figure 39A). The neoplastic cells may be distributed throughout the body or aggregated among normal haemocytes. Blood Sample - Blood samples (smear or suspension) show the characteristics noted for histology. Note - haemocytes with advanced neoplastic changes may be unable to adhere to the microscope slide (Figure 39B, C) so slides should be coated with Poly-L-Lysine.

Prevention - There is no known treatment. It is advisable to avoid introduction of stocks demonstrating haemic neoplasia into areas where neoplasia has not been previously detected. It is also advisable to avoid transfer of stocks from areas where haemic neoplasia causes significant mortalities to areas where no such losse,s have occurred.

References - Brousseau, D.J. 1987. Seasonal aspects of sarcomatous neoplasia in Mya arenaria (soft- shell clam) from Long Island Sound. Journal of Invertebrate Pathology 50:269-276.

Farley, C.A. 1976. Proliferative disorders in bivalve mollusks. Marine Fisheries Review 38(10):30-33.

Farley, C.A., S.V. Otto and C.L. Reinisch. 1986. New occurrence of epizootic sarcoma in Chesapeake Bay soft shell clams Mya arenaria. Fisheries Bulletin 84(4):851-857.

Oprandy, J.J., P.W. Chang, A.D. Pronovost, K.R. Cooper, R.S. Brown and V.J. Yates. 1981. Isolation of a viral agent causing haematopoietic neoplasia in the soft-shell clam Mya arenaria. Journal of Invertebrate Pathology 38:45-51. (continued on page 88) I

I 86 I I I I I a t . . ^^ ^s • .^; ^ ^ ,[^ -^^ ^ir,.^ ^^^ . •^ ^ aa , /1 ^ v,

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Figure 39: (A) Cross-section through the digestive gland of a soft-shell clam (Mya arenaria) I with haemic neoplasia. Note the characteristic enlarged and diffuse nuclei and corresponding reduction of cytoplasm in neoplastic cells (n) compared with normal haemocytes (h). Mitotic figures (m) are also present. Compare with haemic neoplasia of blue mussels (Figure 30B). I (H&E, Scale bar = 100 µm). I I I I

-.•, •e •-•• • • -•;• -••- • ••;•• 11 :6,4 •• w•a' le t • •••••4• • • ••• -• -• • - 1 •• • S&P • • • • b w . • . • • ■ fb•- ••• • • • • 10_1 ., 0 e• • • w • ■ • • ••• • • • ' A w_ w • • • du e .• - . - „. .., • • • •••, •• , ••; ■ e e • e . -•-•• • • • • a' ••• • ••••• ••,:• • • b.; • • e ge.,, .4, . • . • • • • 9..• • •• 14.,••• • • • :to be , • • *s lit • 4111410 lie • • • * P • • . • epe • e• e • • • • ••• • •• • • • • ,/•• • •• • ,•e • • . • • ger:.• n &Nab • • • • • . . ‘ . * few ,•••• t • ••• • 11- • • • • ■ • # • lb: ii• • / ...teed 'fee 6 • ‘e• • ••• s • • • • • •• c • %. . • , • • • • •• • • • g • * * • • • • • _••••ta a • • ..ogee • • • NIP • ••• • •4,-.: -me. .. • •• •ea111 • • .. • 1•• a • • • •:•11•• ••• • • • • •ar '. ..n • a •4 ••_ •••

- w qv - ••• .• if.w•-• ••-i• • • ••-•.••• . • • .- •,-; , .0 ,••• gee •.- • I • • • ( * ■ • . •• f._• ••• •••• V.-• • • 1, • es ••• • e • ••• • . ••• • 1" • • •• it -9 4h. • •• • • -. • • e . 6• • - yr •• • . • • •• •' • le • ' ••* ••• leeie•• •., • • • • .■ 1•• • .• • • w .. ...,• 400 ...••L . • •• eir••• ,' • • se- le' owe IIP• i• ,_ ea • e-% or _ • • limbs so • . 6 .., ,, • •• -• • •• • • • • •• deibw 67 • • • , es*. • ••.6•• er e l.. • • ••••..e • • • • • • 6 - . . •ir•àr. • • • • • . 90 0 9 •• • •el• * f ee•, 9 i b ••• .• fa • •_ «9 • _ • at a_ • .... 9_, _.• IP_ _ • ' Figure 39: (B) Haemocyte suspension preparation from a healthy soft-shell clam (M arenaria). Note the pseudopodial extensions of the normal haemocytes and small, granular nuclei. (Feulgen Picromethyl Blue (FPM), Scale bar = 20 gm). (C) Normal (n) and neoplastic (c) haemocytes in a haemolymph sample from a soft-shell clam (M arenaria) with advanced haemic neoplasia. Note that most of the neoplastic cells are rounded up and show no pseudopodial extensions of the cytoplasm. (FPM, Scale bar = 20 gm). 88

Digenean metacercariae

Background Information - Unidentified Echinostomatid and Gymnophallid metacercariae (larvae) occur in clams (Mya arenaria and Mercenaria mercenaria) throughout Atlantic Canada. They encyst in the connective tissues of the mantle, foot and digestive gland or, more rarely, in the gills. The cyst may develop into a pearl which makes identification of the digenean difficult. In clams from Atlantic Canada infections are usually low (1 per tissue section).

Effect on Host - Clams control penetration by metacercariae by enveloping them in granulomatous lesions which may then develop into pearls. Despite tissue disruption directly around the metacercariae, surrounding tissues appear unaffected and the effect on the host is localised.

Commercial Significance - Pearls in the soft-tissues may reduce clam marketability (see page 76).

Diagnosis - Gross Observation: Observation of pearls of various sizes on the mantle indicates the possible presence of digenean metacercariae. Note -pearls may be initiated by any irritant too large to be expelled by host haemocytes. Histology: Metacercariae occur in the connective tissue of the mantle, gills and digestive gland where they are usually surrounded by intense haemocyte infiltration and an eosinophilic (pink-staining) capsule (the pearl). Metacercariae can be differentiated from other foreign objects stimulating pearl-formation by the presence of body-parts (including suckers, spiny integument, etc) or a distinct vacuole in the centre of the pearl (Figure 40). Note - Pearls may impede sectioning of tissue-blocks. Chipping of microtome blades and consistent tears along the ribbon indicate calcareous inclusions in the tissues. Heavily infected tissues can be fixed in an acidic fixative (e.g. Davidson's) and/or declacified in EDTA (ethylenediaminetetraacetate) post-fixation and prior to paraffin embedding.

Prevention - The,se parasites do not present a significant health problem to clams and, since prevalences are generally low (less than 3%), do not appear to require prevention.

References - Lauckner, G. 1983. Diseases of mollusca: Bivalvia. pp. 632-762. In: ICinne, O. (ed) Diseases of Marine Animals, Vol II. Biologische Anstalt Helgoland, Hamburg.

(continued on page 90) I I 89 I I I I I I I I I I

Figure 40: Echinostome metacercaria (m) encysted within a pearl (p) in the gills of a I quahaug (M. mercenaria). Note tiny spines (sp) around the anterior end of the body. (H&E, I Scale bar = 100 µm). I I I I 90

Odostomia spp ("clam siphon snail")

Background Information - Pyramidellid Gastropods Odostomia seminuda and O. trifida and O. bisuturalis, also known as "clam siphon snails", are found on soft-shell clams (Mya arenaria) from the Gulf of St. Lawrence and Bay of Fundy. The snails are small and white and attach to the openings of the clam siphons. Similar snails (Boonea (= Odostomia) impressa) have been reported on the shells of oysters (Crassostreavirginica and Ostrea edulis) from the United States (Allen 1958) but none have been found, to date, on oysters from Atlantic Canada.

Effect on Host - Siphon snails may have an irritant effect interfering with normal feeding and siphon activity, especially where present in large numbers. The snails use a proboscis to penetrate the hosts' tissues and some species, which live on edible oysters (Ostrea edulis) in the United States, have been associated with a host-tissue response. B. impressa has been shown to impede oyster feeding, however, no similar effect has been observed for clams infested by Odostomia.

Commercial Significance - None lcnown.

Diagnosis - Gross Observation: White snails less than 3 mm in length with a single suture in the columella. Pre,sent on the tips of soft-shell clam siphons (Figure 41).

Prevention - To date no treatment or management techniques are known. Since numbers of Odostomia spp. found on soft-shell clams in Atlantic Canada, to date, do not appear to be pathogenic, prevention may not be necessary.

References - Allen, J.F. 1958. Feeding habits of two species of Odostomia. Nautilus 72: 11-15.

Medcof, J.C. 1948. A snail commensal with the soft-shelled clam. Journal of the Fisheries Research Board of Canada 7: 219-220.

(continued on page 92) 91

Figure 41: Siphons of soft-shell clam (M. arenaria) infested by the marine snail Odostomia seminuda. The small white snails (<3 mm long) are found clustered around the opening of the inhalant siphon. (Scale bar = 5 mm). I 92

I SCALLOP PARASITES, PESTS AND DISEASES I Perkinsus karlssoni Background Information - Perkinsus karlssoni (Protozoa: Apicomplexa: Perkinsidae) was introduced with bay I scallops (Argopecten irradians) into Atlantic Canada in the early 1980's and passed from generation to generation until detected in the late 1980's in hatchery broodstock. Although difficult to detect in developing juveniles, P. karlssoni becomes evident as the bay scallops I mature (towards the end of the first summer of growth). P. karlssoni is believed to pass from infected broodstock to their offspring during spawning and has not been transmitted to native bivalves under close proximity holding experiments. Infective biflagellate spores I can be observed among D-stage scallop larvae within 48 hours of spawning. The parasite has been found in both populations of bay scallops introduced into Atlantic Canada (1979 I and 1989) and occurs in wild populations along the eastern United States. Effect on Host - Since bay scallops do not normally survive long after spawning, the specific pathogenicity I of the parasite is difficult to assess. Based on tissue damage around the mantle, where the parasites aggregate in advanced infections, it is assumed that they may play a secondary role in reducing the bay scallop's ability to recover post-spawning. Related species found I in the clam Ruditapes decussatus, as well as in Crassostrea virginica from the eastern US and abalone (Haliotis spp) from Australia, are all associated with host mortality. I Commercial Significance - Although the pathogenic significance of P. karlssoni is difficult to assess, it does not appear to be a significant problem for the bay scallop or other bivalve species.

I Diagnosis - Iiistology: Observation of eosinophilic protozoans with densely basophilic inclusion bodies throughout the tissues (Compare Figure 42A, B, with Figures 43A, B). Sizes of 1 parasite lesions range from 12 µm (individuals) to over 75 µm (multiplicative stages). Proliferation of P. karlssoni is preceded by a massive infiltration of haemocytes throughout the connective tissue of the digestive gland, mantle and gonad. The optimum I time for detection is during or after spawning, when P. karlssoni spreads throughout the tissues (especially the mantle). Extensive deposition of ceroid (similar to that described for Perkinsus marinus ("dermo") infections of eastern oysters, Crassostrea virginica) makes I identification of the parasite difficult in advanced infections. Thioglycollate Culture: Appearance of blue-black Lugol-positive spheres in the tissue of bay scallops which have been incubated in fluid thioglycollate broth for at least 7-10 days I at room temperature (> 20°C) (Figure 43C). Prevention - I Since no mass mortalities of bay scallops have been directly attributed to Perkinsus I karlssoni, and it does not infect other bivalve species, control appears unnecessary. I I 93

References - Karlsson, J.D. 1991. Parasites of the bay scallop, Argopecten irradians (Lamarck, 1819). pp 180-190. In: Shumway, S.E. and P.A. Sandifer (eds) An International Compendium of Scallop Biology and Culture. World Aquaculture Workshops, Number 1, World Aquaculture Society and National Shellfisheries Association, Baton Rouge, LA..

McGladdery, S.E., R.J. Cawthorn and B.C. Bradford. 1991. Perkinsus karlssoni n. sp. (Apicomplexa) in bay scallops Argopecten irradians. Diseases of Aquatic Organisms 10:127-137.

McGladdery, S.E., B.C. Bradford and D.J. Scarratt. 1993. Investigations on the transmission of parasites of bay scallops (Argopecten irradians) introduced to Canadian Atlantic waters. Journal of Shellfish Research 12(1):49-58.

Whyte, S.K., R.J. Cawthorn and S.E. McGladdery. (in press) A comparison of the pathology caused by Perkinsus karlssoni (Apic,omplexa, Perkinsea) and an unidentified coccidian parasite (Apicomplexa) in the bay scallop Argopecten irradians. Disease,s of Aquatic Organisms.

(continued on page 98) 94

Figure 42: (A) Haematoxylin and eosin stained tissue-section of a healthy bay scallop (Argopecten irradians). Mantle (m), female gonad (fg), male gonad (mg), gills (gl), digestive gland (dg), stomach (s), intestine (i) and kidney (k). (Scale bar = 2 mm). I

g I

Figure 42: (B) Haematoxylin and eosin stained tissue-sections of a healthy giant sea scallop (Placopecten magellanicus). Tissue sections of whole scallops >75 mm wide will not fit on a single microscope slide (as shown in Figure 42A). Separate samples are taken through the digestive gland (dg) and kidney (k), as well as the gills and gonad (Figure 42C). (Scale bar = 2 mm). (C) Haematoxylin and eosin stained tissue-sections through the gills (gl) and gonad (gd) of a giant sea scallop (Placopecten magellanicus). (Scale bar = 2 mm). 96

Figure 43: (A) Cross-section of bay scallop, Argopecten irradians, showing "swirl" lesions induced by Perkinsus karlssoni (H&E, Scale bar = 100 gm). (B) High-power magnification of P. karlssoni within an early host haemocyte encapsulation response. Note flattened haemocytes (h) around the parasites, which are characterised by the presence of vacuoles and dense inclusion bodies (ib). (H&E, Scale bar = 25 gm). I

I 97 I I I I I I I I I I

Figure 43: (C) Lugol-positive spores of P. karlssoni, grown in thioglycollate medium from I infected bay scallop (A. irradians) mantle tissue (Scale bar = 100 gm). I I I I I I 1 I 98 Kidney Coccidia I Background Information - Kidney coccidians, including various species in the genus Pseudoklossia, are found in a number of bivalve species. The identity of the kidney coccidian found in bay scallop I (Argopecten irradians) appears to be a new species of Pseudoklossia (Dr. R.J. Cawthorn, pers. comm.) and its life-cycle is currently under investigation. It appears to proliferate under warm water conditions (temperatures > 20°C) and has passed from generation to generation from bay scallops introduced to Canada in thé late 1979-80. This indicates I that, like P.karlssoni (page 92), the kidney coccidian may be transmitted from infected broodstock to their offspring during spawning. So far, no similar speçies have been detected in other bivalve species in Atlantic Canada. The same kidney coccidian is known to infect bay scallops along the eastern United States.

Effect on Host - I Infection by the bay scallop pseudoklossian is not generally associated with a specific host-response and no mortalities have been attributed to the parasite in wild populations from the eastern United States. A single case of over 90% mortality, however, has been 1 reported from Atlantic Canada (Cawthorn et al., 1992). This occurred following transfer of infected bay scallops from a shellfish hatchery to experimental holding facilities using warm (> 20°C) recirculating artificial seawater. The coccidian was found throughout the I soft-tissues of moribund bay scallops. Juvenile bay scallops from a US hatchery have also been reported as hosting pathogenic infections. Similar coccidians have been reported to cause mortalities of sea scallops (Pecten maximus) in France. Pathogenicity appears to be I related to the spread of the parasite from the kidney to the other tissues. Commercial Significance - I The bay scallop pseudoklossian is not normally considered a significant health concern. However, under warm-water holding conditions the parasite can proliferate to intensities which are significantly pathogenic.

I Diagnosis - Histology: Presence of eosinophilic protozoans in the epithelial cells of the kidney or loose within the renal tubules (Figure 44A). Infected cells may or may not be I hypertrophied. In severe infections these coccidians may be found in the epithelium of the stomach or intestine (Figure 44B) and sizes range from 6 µ.m to 30 µm. The parasites have granular cytoplasm, a single spherical nucleus and vary in shape from spherical to I oblong or pear-shaped. Several life-history stages can be seen in the same host.

Prevention - i All mortalities reported to date are correlated to adverse or "stressful" conditions although pathogenicity may also be related to the species of Pseudoklossia-like protozoan. Under hatchery conditions pathogenicity can be controlled by reducing stocking density and I lowering water temperatures (less than 20°C). The use of artificial seawater has also been suggested as a contributing factor (Cawthorn, R.J., pers.comm.). Since kidney coccidians have a ubiquitous distribution in a wide variety of bivalve species, prevention I may be impractical. I I 99

References - Cawthorn, R.J., R.J. MacMillan and S.E. McGladdery. 1992. Epidemic of Pseudoklossia sp. (Apicomplexa) in bay scallops Argopecten irradians maintained in a warm water recirculating facility. Canadian Society of Zoologists Meeting, Antigonish, N.S., May, 1992 (Abstract only).

Getchell, R.G. 1991. Diseases and parasites of scallops. pp 471-494. In: Shumway, S.E. (ed.) Scallops: Biology, Ecology and Aquaculture. Developments in Aquaculture and Fisheries Science #21. Elsevier.

Karlsson, J.D. 1991. Parasites of the Bay Scallop, Argopecten irradians (Lamarck, 1819). pp 180-190. In: Shumway, S.E. and P.A. Sandifer (eds.) An International Compendium of Scallop Biology and Culture. World Aquaculture Society Workshops, Number 1, World Aquaculture Society, Baton Rouge, LA., 357pp.

McGladdery, S.E. 1990. Shellfish parasites and diseases on the east coast of Canada. Aquaculture Association of Canada Bulletin 90-3: 14-18.

Whyte, S.K., R.J. Cawthorn and S.E. McGladdery. (in press) A comparison of the pathology caused by Perkinsus karlssoni (Apicomplexa, Perkinsea) and an unidentified coccidian parasite (Apicomplexa) in the bay scallop Argopecten irradians. Diseases of Aquatic Organisms.

(continued on page 101) • 100

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Figure 44: (A) Bay scallop, A. irradians, lcidney infected by Pseudoklossia-like coccidian parasites (p). (rtl - renal tubule lumen; ke - lcidney epithelium). (H&E, Scale bar = 100 Inn). (B) Pseudoklossia-like coccidians (p) between the epithelial cells of a bay scallop, A. irradians, stomach (es). (dt - digestive tubules, ct -connective tissue infiltrated by haemocytes). (H&E, Scale bar = 100 kun). 101 Digeneans

Background Information - Digenean larvae are found in a wide range of bivalve species worldwide including bay scallops (Argopecten irradians) and giant sea scallops (Placopecten magellanicus) from Atlantic Canada. They are found either as encapsulated metacercariae (late larval stage) or, more rarely, unencapsulated in the connective tissue. The identity of encapsulated digeneans is difficult to determine from tissue sections but, base,d on the presence of spines around the anterior sucker, most appear to be echinostomatid or gymnophallid digeneans. They usually occur in low numbers (1-2) per tissue-section.

Effect on Host - Metacercarial infections stimulate haemocyte infiltration and encapsulation, which may develop into a pearl (Figure 45), however there is no evidence of extensive tissue damage. Metacercariae lying free in the tissues (Figure 46) show evidence of tissue disruption around the point of penetration but haemocyte infiltration is focal.

Commercial Significance - High numbers which could affect marketability of whole scallops have not been found to date. Meats are generally not affected.

Diagnosis - Gross Observation: Pearls indicates the possible presence of digenean metacercariae. Note - pearls may be initiated by any irritant too large to be expelled by host haemocytes. Histology: Metacercariae are found in the connective tissue of the mantle, gills, digestive gland and, rarely, the kidney (Figure 45). They are surrounded by haemocytes and/or an eosinophilic (pink-staining) capsule which indicates calcium carbonate (pearl) deposition. Metacercariae can be differentiated from other foreign objects where body-parts, such as suckers or integument, are visible inside the pearl. Unencapsulated digeneans are usually found in the connective tissue of the digestive gland (Figure 46) and can be distinguished from turbellarians (page 57 and 109), by surface striations and/or spines, as well as two suckers. Note - Pearls may be impede sectioning of tissue-blocks. Heavily infected tissues can be fixed in an acidic fixative (e.g. Davidson's) and/or declacified in EDTA (ethylenediaminetetraacetate) post-fixation and prior to paraffin embedding.

Prevention - These parasites do not appear to require preventative measures.

References - Lauckner, G. 1983. Diseases of mollusca: Bivalvia. pp. 632-762. In: Kinne, O. (ed) Diseases of Marine Animals, Vol II. Biologische Anstalt Helgoland, Hamburg. (review of literature on the effects of all stages of digeneans on various marine bivalve species).

McGladdery, S.E. and M.F. Stephenson (unpublished data). Atlantic Canadian Association of Parasitologists Meeting, Mont-Joli, Quebec, October 3-4, 1991.

Stunkard, H.W. 1938. The morphology and life-cycle of Himasthla quissentensis (Miller and Northrup, 1926). Biological Bulletin of the Marine Biology Laboratory (Woods Hole)75: 145-164. (continued on page 103) 102

Figure 45: Echinostome metacercaria encysted within the kidney of a bay scallop, A. irradians. Note haemocyte encapsulation (ec) and spiny integtunent around the anterior end of the parasite. N.B. there is no evidence of pearl encapsulation (see Figures 35B, C, 40). (H&E, Scale bar = 100 pm).

Figure 46: Unidentified digenean in the digestive gland of giant sea scallop, P. magellanicus. Note unciliated, serrated integument (st), oral sucker (os) and haemocyte infiltration (h) into the connective tissue in front of the parasite. (H&E, Scale bar = 100 gm). I I 103 Bacterial Abscess Lesions (Brown Spot Disease) I Background Information - "Brown-spot", or bacterial abscess disease, has been reported in giant sea scallops (Placopecten magellanicus) from Atlantic Canada and the eastern United States. It is I easily visible with 1-4 mm wide brown-spots in the white meats. It usually occurs in larger scallops (> 100 mm shell height) and the cause has yet to be clearly identified. Various bacteria have been cultured from "brown spot" lesions. Getchell (1991) described i a Gram-positive pleomorphic bacterium from populations of giant sea scallop in Maine. Gram-negative Vibrio and Pseudomonas species have been isolated from lesions in scallops from the Gulf of St. Lawrence. In most cases, the infection is spread throughout the soft-tissues. Some kind of irritation may be responsible for initiating ulceration and I subsequent infection, however, shell fouling and boring have not been consistent features of cases reported so far, although most appear localised in estuarine or in-shore localities. The distribution of brown-spot disease is patchy with unaffected scallops being found I within 100 m of highly infected beds. The causative agent does not appear to be highly contagious. Prevalence of "brown spot" ranges from 4-12% and have only been found, to I date, in bottom-grown scallops. Effect on Host - Abscessing and necrosis of the muscle fibres may affect swimming and closing capability, I however, there is no direct evidence of mortality due to this disease. Commercial Significance - I Meats affected by "brown spot" are unmarketable and are disposed of during shucking. Diagnosis - I Gross Observation: Presence of brown pustules, ranging in diameter from 1-4 mm, embedded in the adductor muscle (Figure 47A). Histology: Presence of abscesses containing necrotic host tissue, haemocytes and I bacteria. Lesions are usually largest in the adductor muscle (Figure 47B), but focal haemocyte aggregations may also be found in the digestive gland, kidney and gonad. Bacteria are not always evident.

I Prevention - There are no known methods to control or prevent this problem.

I References - Getchell, R.G. 1991. Diseases and parasites of scallops. pp 471-494 In: Shumway, S.E. (ed.) Scallops: Biology, Ecology and Aquaculture. Developments in Aquaculture and I Fisheries Science #21, Elsevier.

McGladdery, S.E. 1990. Shellfish parasites and diseases on the east coast of Canada. I Aquaculture Association of Canada Bulletin 90-3: 14-18. i (continued on page 105) I I • ▪▪

104

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Figure 47: (A) "Brown-spot" lesion (arrow) in the adductor muscle of a giant sea scallop, Placopecten magellanicus (Scale bar = 1 mm). (B) Cross-section through muscle fibres (ml) of a giant sea scallop, P. magellanicus, affected by "brown spot". The abscess lesions are filled with necrotic haemocytes (nh). (H&E, Scale bar = 100 gm). I 105

1 T3richodina-like Gill Ciliates

Background information - I Trichodinid ciliates are found on the surface of giant sea scallops (Placopecten magellanicus) from Atlantic Canada. They occur on the gills but may also be found on the surface of the kidney and gonad. Similar trichodinids are reported from Japanese I scallop (Patinopecten yessoensis) and species of Chlamys. The identity of the trichodinid on giant sea scallop is currently being studied. Numbers increase over the summer peaking (-60-95% prevalence and >20 per tissue-section) in early October. Infection I levels over winter are unknown. Interestingly a comparison between wild bottom-growing populations and scallops grown in suspension showed that both host these ciliates, I although the bottom-grown stocks had higher prevalences and intensities of infection. Effect on Host - Numbers of trichodinids are usually low (< 10 per gill-section), however, over 70 have I been seen on a single tissue section. No obvious host-reponse has been observed to these infections. This is similar to the lack of host response noted for the other trichodinids from Japanese scallop (Patinopecten yessoensis) and Chlamys spp. from the I Commonwealth of Independent States. Commercial Significance - I None known. Diagnosis - Gill Smear: Presence of "flying-saucer"-like ciliates characterised by a ring of denticles 1 and horse-shoe shaped macronucleus. Lateral views of living specimens look like English bowler hats. r Histology: Presence of ciliates, as described above, on or in close association with the gill filaments (Figure 48A, B). The ring of denticles may persist after the rest of the ciliate has been destroyed (e.g., when processing scallops which have been out of I seawater for over 12 hours). Prevention - Since numbers are usually low and there has been no evidence of pathogenicity to date, I prevention appears unnecessary.

References - I Beninger, P.G., M. LePennec and M. Salaun. 1988. New observations of the gills of Placopecten magellanicus (Mollusca: Bivalvia), and implications for nutrition. Marine I Biology, 98: 61-70. Getchell, R.G. 1991. Diseases and parasites of scallops. pp 471-494 In: Shumway, S.E. (ed.) Scallops: Biology, Ecology and Aquaculture. Developments in Aquaculture and I Fisheries Science #21. Elsevier. McGladdery, S.E. 1990. Shellfish parasites and diseases on the east coast of Canada. I Aquaculture Association of Canada Bulletin 90-3: 14-18. (continued on page 107) I I

I 106 I I I I I I I

I B i 1 I I I

.1: I I Figure 48: (A) Cross-section of giant sea scallop, P. magellanicus, gills with Tri chodina-like ciliates (arrows) (H&E, Scale bar = 100 gm). (B) Trichodinid-like ciliate on the gills of I giant sea scallop, P. magellanicus (H&E, Scale bar = 20 gm). I I I I 107 Gill Turbellaria

Background Information - I Unidentified turbellarians (Platyhelminthes: Turbellaria) have been found occasionally on giant sea scallop (Placopecten magellanicus) from the Gulf of St. Lawrence. Numbers are usually low (< 5% prevalence and 1-2 per gill-section) and whole specimens have not I yet been collected for specific identification. Scallop turbellarians resemble Urastoma cyprinae from eastern oysters (Crassostrea virginica) (see page 56). Gill turbellarians are I found both suspension- and bottom-grown giant sea scallops. Effect on Host - r No host-response has been observed. Commercial Significance - I None known. Diagnosis - Gross observation: Prevalence has been so low (< 5%) that whole specimens have yet to I be collected. Histology: gill and palp sections showing ciliated flatworms (Figure 49A, B). I Prevention - Since numbers rarely exceed 1-2 per gill-section and there is no obvious tissue damage, I prevention appears unnecessary. References - I McGladdery, S.E. and M.F. Stephenson. Unpublished data. Stafford, D. 1912. On the Fauna of the Atlantic Coast, Third Report. Contributions to I Canadian Biology 1906-1910, pp 45-67., C.E. Parmelee, Ottawa 305p. I I I I I I (continued on page 109) I 108

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_ C•-•

" 42' e

F..- • — re in N.sa ou • _ e, 7." f■L - C

•••■••

Figure 49: (A) Cross-section of giant sea scallop, P. magellanicus, gills and a surface turbellarian (H&E, Scale bar = 0.5 mm). (B) Turbellarian from the gills of giant sea scallop, P. magellanicus magnified to show ciliated epidemis (H&E, Scale bar = 100 gm). 109

Intestinal Turbellaria

Background Information - Unidentified turbellarians (Platyhelminthes: Turbellaria) are occasionally found in the intestine of giant sea scallops (Placopecten magellanicus) from the Gulf of St. Lawrence. Prevalence is low (less than 1%) and only 1 per tissue-section has been observed, to date. Whole specimens have yet to be collected in order to identify the,se turbellarians to species. No turbellarians have been reported from scallop species elsewhere.

Effect on Host - No tissue damage or host-response to these turbellarians has been detected.

Commercial Significance - None known.

Diagnosis - Histology: Presence of ciliated worms (Figure 50) in sections through the digestive gland of giant sea scallops.

Prevention - Since turbellarians have only been found occasionally in the intestines of giant sea scallops and are not associated with pathology, prevention appears to be unnecessary.

References - McGladdery, S.E. and M.F. Stephenson. Unpublished data.

(c,ontinued on page 111) I I 110 I I I I I I I I I I

Figure 50: Cross-section through the digestive gland of a giant sea scallop, P. magellanicus, I containing a turbellarian (tb). Note ciliated epidermis (ce) and pharynx (ph) (H&E, Scale bar I 100 µm). I I I I I I 111

I ACKNOWLEDGEMENTS

This manual could not have been compiled without the generous support and assistance of the I following people: Mr. Arnold Ross, Mr. Roger Townsend and Mr. Wayne Somers (Prince Edward Island); Mr. Gerald Cormier, Messrs. Serge and Reginald Dugas and Mr. Maurice Daigle (New Brunswick); SFT Ventures and Messrs. John Harding and Peter Darnell (Nova I Scotia) and the Port-au-Port Development Association (Newfoundland). Many other producers, too numerous to mention, also provided valuable material for this manual.

I Special thanks are also given to all the respective Provincial Departments of Fisheries and Aquaculture in Atlantic Canada, as well as DFO Gulf Region Area Offices, for continuous advice and support of this survey. Dr. Susan Bower, DFO, Pacific Biological Station, and Drs. Carol I Morrison and David Scarratt and Mr. John Cornick, DFO Halifax have also been sources of regular and valuable advice. Mr. R.E. Drinnan provided welcome advice and constructive criticism throughout the compilation of this manual, as well as invaluable information from his I mountainous collection of shellfish data and material. We (McGladdery and Stephenson) were honoured when he accepted our invitation to be a co-author. Valuable material was also provided by Dr. David Alderman, M.A.F.F., Fish Diseases Laboratory, Weymouth, England and Dr. I M.F. Li from their research on Malpeque Disease and Shell Disease. We gratefully acknowledge the support of our Molluscan Section colleagues and summer students, who collected and helped process many of our samples. Special thanks are also due to Dr. T.W. Sephton, Molluscan I Section.Head, for his reviews and consistent support for the manual and its colour reproduction. Mr. Robert Maillet, Photographer, Moncton, produced the shell and whole body photography (black background) for this manual and demonstrated incredible patience in making what we I needed to point out clearly visible. Anyone wishing a colour print reproduction of any of these photographs should address their request directly to him. The illustrations indicating the source of tissue sections throughout this manual were modified from the excellent diagrams produced by I Alice Jane Lippson for the NOAA Technical Memorandum NMFS-F/NEC-25 "Histological Techniques for Marine Bivalve Mollusks", Howard, D.W. and C.S. Smith (1983). Her work has greatly helped us both for this manual and for teaching bivalve pathology at various institutions I throughout the Maritimes.

Last, but by no means least, we thank Mr. C. Austin Farley of the Cooperative NOAA I Laboratory, Oxford, Maryland, for sharing his boundless knowledge and `infectious' enthusiasm I for bivalve parasites, pests and diseases. I I I I I 112

LIST OF FIGURES

EXAMPLES OF TERMS COMMONLY USED IN TEXT

Figure lA Multiple granulomas (g) in the connective tissue of bottom-grown blue mussels (M. edulis). (dt - digestive tubules, int - intestine) (H&E, Scale bar = 0.5 mm) (P. 3). Figure 1B Granuloma abscess lesion containing eosinophilic (pink) granulocytes, necrotic cells and cell debris. (H&E, Scale bar = 100 gm). (p. 3).

Figure 2A Eosinophilic (pink-staining) adipogranular cells in the connective tissue of blue mussels (M. edulis) - used for storage of energy sources (glycogen, lipids etc.). (dt - digestive tubules, mg - male gonads) (H&E, Scale bar = 100 gm). (p. 4).

Figure 2B Blue mussel (M. edulis) with no evidence of stored energy (i.e., adipogranular cells "empty"). (dt - digestive tubule, mg - male gonads) (H&E, Scale bar = 100 gm). (p. 4).

Figure 3A Ceroid-containing ("brown" or cerous) cells (arrows) spread through the connective tissue of eastern oysters (C. virginica). (H&E, Scale bar = 0.5 mm). (P. 5). Figure 3B Ceroid-containing cells demonstrating "colour" and size variation. (H&E, Scale bar = 100 gm). (p. 5).

Figure 4 Concretions (c) within the epithelial cells of the digestive tubules (dt) of giant sea scallop (P. magellanicus). (H&E, Scale bar = 100 gm). (p. 6).

Figure 5 Diapedesis - haemocyte (h) migration across epithelial walls- as shown here across the intestinal epithelium (e) of an eastern oyster (C. virginica). Note accumulation of haemocytes within the lumen (1) of the intestine. (H&E, Scale bar = 100 gm). (p. 6).

Figure 6 Duct (d) and tubule (t) epithelial cells in the digestive gland of an eastern oyster (C. virginica). (H&E, Scale bar = 100 gm). (p. 7).

Figure 7A Ripe male gonads of eastern oyster (C. virginica) showing sperm (s) cells ready for release via the gonoducts (gd). (H&E, Scale bar = 100 gm). (p. 9).

Figure 7B Ripe female gonads of eastern oyster (C. virginica) showing eggs at various stages of development from early (e) to stalked/pedunculated (p) to ripe (r). (H&E, Scale bar = 100 gm). (p. 9).

Figure 7C Hermaphroditic (both sexes) gonad of edible oyster (Ostrea edulis) showing ripe sperm (s) and ova (o). Note difference between the densely basophilic (blue) sperm and eosinophilic (pink) ova. (II&E, Scale bar = 100 gm). (p. 10). 113

Figure 7D Eastern oyster (C. virginica) with no ganaetes in the gonad. Note haemocyte (h) infiltration into the germinal tissues (gt) and the gonoducts (gd). (H&E, Scale bar = 100 gm). (p. 10).

Figure 7E Post-spawning eastern oyster (C. virginica) showing haemocyte infiltration into the gonads and gonoducts (gd), as well as egg resorption (er). (H&E, Scale bar = 100 gm). (p. 11).

Figure 8A Negligible haemocyte infiltration in an eastern oyster (C. virginica). (connective tissue (et), digestive tubules (dt), artery (a) and intestine (i). (H&E, Scale bar = 100 gm). (p. 12).

Figure 8B Extensive haemocyte infiltration almost completely obscuring the connective tissue. (digestive tubules (dt) and digestive duct (dd)). (H&E, Scale bar = 100 gm). (p. 12).

Figure 8C Localised (focal) infiltration (arrows) in the connective (et) tissue of eastern oysters (C. virginica). -ceroid-containing cells). (H&E, Scale bar = 100 gm). (p. 13).

Figure 9 • Hypertrophy of digestive tubule epithelium (dt) in eastern oyster (C. virginica), infected by a rickettsia-like organism (r). (H&E, Scale bar = 100 gm). (p. 14).

Figure 10 Erosion of the nacreal layer of a giant sea scallop (P. magellanicus) shell due to prolonged mantle recession. (p. 15).

Figure 11A Epithelial cell metaplasia (change in cell shape from columnar to cuboidal or squamous (= flattened)) in the digestive tubules (dt) of a healthy eastern oyster (C. virginica). See also Figure 6. (H&E, Scale bar = 0.5 mm). (p. 17).

Figure 11B Extreme metaplasia in a moribund eastern oyster (C. virginica), possibly due to chronic starvation. All cells of the digestive tubules (dt) are flattened. Note, the digestive ducts (dd) are not changed. Compare with Figure 6. (H&E, Scale bar = 0.5 mm). (p. 17).

Figure 12 Tissue necrosis indicated by connective tissue (et) and epithelial (ep) breakdown and diffuse haemocyte (h) infiltration, in a moribund eastern oyster (C. virginica). (H&E, Scale bar = 0.5 mm). (p. 18).

Figure 13A Haemocyte neoplasia in a blue mussel (M. edulis). Note heavy infiltration throughout the connective tissue and reduced profiles of the digestive tubules (tb) and ducts (dc). (H&E, Scale bar = 0.5 mm). (p. 19).

Figure 13B Neoplastic haemocytes in a blue mussel (M. edulis). Note abnormally large and dense-staining nuclei in relation to reduced cytoplasm, as well as the high prevalence of mitotic figures (mf). (H&E, Scale bar = 100 gm). (p. 19). 114

Figure 14A Surface view of oedema (o) along the dorsal line of the mantle (mt) of eastern oyster (C. virginica). (Scale bar = 5 mm). (p. 20).

Figure 14B Oedema (o) of the mantle connective tissue (et) in a blue mussel (M. edulis). (II&E, Scale bar = 0.5 mm). (p. 20).

GENERAL PARASITES, PESTS AND *DISEASES OF ATLANTIC BIVALVES

Figure 15A Indentation of the shell margin of an eastern oyster (C. virginica). Possibly caused by crowded growing conditions or growth around a foreign object. (p. 22).

Figure 15B Raised "knob" of conchiolin on the inner shell surface of an eastern oyster (C. virginica). (p. 22).

Figure 15C Shell growth around a plastic tag used to hang edible oyster (O. edulis) for suspended culture. (p. 23).

Figure 15D Pearl-like formations attached to inner surface of the shell of a blue mussel (M. edulis). Tissue irritants are walled off from the soft-tissues by the deposition of additional nacre (similar to pearl formation within the soft-tissues, e.g., Figures 16C and 35A, B). (p. 24).

Figure 15E Hole bored through the shell of a blue mussel (M. edulis) by a gastropod predator (probably the moon snail, Lunatia heros). (p. 24).

Figure 16A Deformation of the gills of an eastern oyster (C. virginica). Note the presence of several gill turbellarians (arrows) (Urastoma cyprinae) (see also Figure 25). (Scale bar = 5 mm). (p. 26).

Figure 16B Discolouration of the mantle tissues of blue mussel (M. edulis) due to the presence of green algae. (Scale bar = 5 mm). (p. 26).

Figure 16C Pearls in the mantle tissues of blue mussel (M. edulis). Pearls of this size are commonly found in bottom-growing mussels, but rarely in suspension-grown mussels. (Scale bar = 5 mm). (p. 27).

Figure 16D The nemertean clam worm, Cerebratulus lacteus, which lives on soft-shell clams (Mya arenaria). (Scale bar = 1 cm). (p. 28).

Figure 17A Characteristic tunnels of Polydora spp. in the shell of an eastern oyster (C. virginica). (p. 30).

115

Figure 17B Mud blisters on inner shell surface of an eastern oyster (C. virginica), caused by Polydora spp. tunnels. (p. 30).

Figure 17C U-shaped discolourations on the imier surface of a giant sea scallop, P. magellanicus, shell. Tunnel shape varies with the species of Polydora inhabiting the shell. In this case tunnel shape is indicative of P. websteri. (p. 31).

Figure 17D Polydora websteri in the mantle fluids of an eastern oyster, C. virginica. (Scale bar = 5 mm). (p. 31).

Figure 17E Nereis diversicolor from the shell surface of an eastern oyster, C. virginica. (Scale bar = 5 mm). (p. 32).

Figure 17F Serrated "jaws" which characterize Nereis spp. from the shell-boring Polydora spp. (Scale bar = 1 mm). (p. 32).

Figure 18A Surface of an eastern oyster (C. virginica) shell severely excavated by a boring sponge (Cliona spp). (p. 35).

Figure 18B Outer shell surface of a blue mussel (M. edulis) affected by the boring sponges, - Cliona spp. (p. 35).

Figure 18C Outer shell surface of a giant sea scallop (P. magellanicus) affected by the.boring sponges, Mona spp. (p. 36).

Figure 18D Profile of advanced clionid sponge damage throughout the matrix of an e,astern oyster (C. virginica) shell. Note the thickened profile of the shell, which develops as the oyster attempts to prevent perforation through to the inner surface. (p. 37).

Figure 18E Inner surface of an eastern oyster (C. virginica) shell extensively penetrated by the boring sponge Cliona sp. Note raised knobs of conchiolin where the adductor muscle attaches (dark annulated scar area). (p. 37).

Figure 18F limer surface of a blue mussel (M. edulis) shell perforated by clionid sponges (Cliona sp). (p. 38).

Figure 18G Inner shell surface of a giant sea scallop (P. magellanicus) perforated by a clionid sponge. Note the brown spots characteristic of sponge penetration, pearl-like formations where the adductor muscle attaches, as well as extensive invasion of the Cliona tunnels by an alga. (p. 38).

Figure 19A "Blue body" (bb) lesions in the haemocytes (h) and sloughed off epithelial cells (ec) of the digestive tubules of an eastern oyster (C. virginica). (H&E, Scale bar = 100 gm). (p. 41).

Figure 19B "Blue body" lesions in the epithelial cells of the digestive tubules of an eastern oyster (C. virginica). (H&E, Scale bar = 100 gm). (p. 41). 116

Figure 19C Cross-section through digestive tubule epithelia (te) of a blue mussel (M. edulis) containing "blue bodies" (bb). (H&E, Scale bar = 100 gm). (p. 42).

Figure 19D Cross-section through gill epithelia (ge) of a blue mussel (M. edulis) with a "blue body" (bb) inclusion (H&E, Scale bar = 100 im). (p. 42).

Figure 19E "Blue body" (arrows) inclusions in the digestive ducts of a quahaug (hard-shell clam) (M. mercenaria). (H&E, Scale bar = 100 gm). (p. 43).

Figure 19F "Blue body" (bb) inclusion in the digestive gland of a bay scallop (A. irradians) (H&E, Scale bar = 100 gm). (p. 44). •

Figure 19G "Blue body" (bb) inclusion in the gill epithelia (ge) of a bay scallop (A. irradians) (H&E, Scale bar = 100 gm). (p. 44).

OYSTER PARASITES, PESTS AND DISEASES

Figure 20 Haematoxylin and eosin stained tissue-section from a healthy eastern oyster (C. virginica). Mantle (m), gonad (gd), gills (gl), digestive gland (dg), stomach (s), • intestine (i) and kidney (k). (Scale bar = 2 mm). (p. 47).

Figure 21A Section through an eastern oyster (C. virginica) with an advanced case of - Malpeque Disease. Note the massive abscess lesions (a) and lack of gonad development. (H&E) (Material provided by Mr. R.E. Drinnan). (Scale bar = 2 mm). (p. 48).

Figure 21B Blood cell proliferation associated with Malpeque disease in an eastern oyster (C. virginica). Note the diffuse, enlarged nuclei (compare with Figure 8B), breakdown of digestive tubule (dt) and digestive duct (dd) epithelia, as well as extensive diapedesis (dp). (Material provided by Mr. R.E. Drinnan). (H&E, Scale bar = 100 gm). (p. 48).

Figure 22A Cross-section through the gills of an eastern oyster (C. virginica) showing Sphenophrya-like ciliates (arrows). (H&E, Scale bar = 0.5 mm). (p. 50).

Figure 22B High magnification of Sphenophrya-like ciliates on the gills of an eastern oysters (C. virginica). Note direct attachment to the gill epithelial cells (arrows) and characteristic dense-staining nuclei (compared with the oyster-tissue nuclei). (H&E, Scale bar = 20 gm). (p. 50).

Figure 23A Cross-section through the digestive gland of an eastern oyster (C. virginica) showing Ancistrocoma-like ciliates (arrows) inside the digestive tubules (dt). (H&E, Scale bar = 100 gm). (p. 52).

Figure 23B Individual Ancistrocoma-like ciliate embedded inbetween the tubule epithelial cells. Note Large, granular macronucleus and tentacle-like attachment to the basal membrane. (H&E, Scale bar = 20 gm). (p. 52). I 117

I Figure 24A Cross-section through the gonad of a female eastern oyster (C. virginica) showing normal ova and ova affected by viral gametocytic hypertrophy (vg). (H&E, Scale I bar = 100 µm). (p. 54). Figure 24B Cross-section through the gonad of a male eastern oyster (C. virginica) showing normal sperm and sperm affected by viral gametocytic hypertrophy (vg). Note I the various stages of hypertrophy (hs) and rupture (rs) of individual sperm cells. (H&E, Scale bar = 100 µm). (p. 54). I Figure 24C Cross-section through immature gonadal tissue of an eastern oyster (C. virginica) showing normal germ cells (ng) and cells infected by the virus (vg). (H&E, Scale I bar = 100 µm). (p. 55). Figure 25 Histological section of the turbellarian Urastoma cyprinae lying between gill- filaments of an eastern oyster (C. virginica) (See also Figure 16A). (H&E, Scale I bar = 100 µm). (p. 58). Figure 26 Unidentified turbellarian (t) containing larvae (1), in the intestine (i) of an eastern I oyster (C. virginica). (H&E, Scale bar = 100 µm). (p. 58). Figure 27A- Shell damage to an edible oyster (O. edulis), infected by the fungus Ostracoblabe implexa. Note the extensive dark discolouration of the calcareous layer , I underlying the thin nacre layer of the shell, as well as extensive shell "wart" formation (arrows) on and around the adductor muscle attachment area of the shell. The hinge and edge of the shell also show characteristic thickening. I (Material kindly provided by Dr. D. J. Alderman). (p. 60).

Figure 27B Hyphal network of Ostracoblabe implexa cultured from infected edible oyster (O. I edulis) shell. (Scale bar = 50 µm). (Material provided by Mr. R.E. Drinnan). (p. 60).

I Figure 28A Hexamita sp. collected from a culture of necrotic quahaug tissues (M. mercenaria). (Scale bar = 100 µm). (p. 63).

1 Figure 28B Necrotic mantle tissue of an eastern oyster (C. virginica) showing aggregations of Hexamita sp. (arrows) in the connective tissue and in the haemolymph, necrotic haemocytes, as well as connective and epithelial tissue breakdown. (H&E, Scale I bar = 20 µm). (p. 63). I MUSSEL PARASITES, PESTS AND DISEASES Figure 29 Haematoxylin and eosin stained tissue-section through the tissues of a healthy blue I mussel (Mytilus edulis). Mantle (m), gonad (gd), gills (gl), digestive gland (dg), stomach (s), intestine (i), foot-retractor muscles (rm) and foot (f). (Scale bar = 2 I mm). (p. 66). I I I 118

I Figure 30A Histological section through a blue mussel (M.edulis) with haemic neoplasia. Note the intense haemocyte infiltration throughout the connective tissue (ct) and the dark staining properties of the affected haemocytes. (Compare with Figure I lA). (H&E, Scale bar = 0.5 mm). (p. 67). Figure 30B Neoplastic haemocytes of a blue mussel (M. edulis) showing enlarged nuclei and I diffuse nucleoplasm, reduced cytoplasm and several mitotic figures (arrows). Some normal haemocytes (n) are present among the neoplastic cells. (dt - I digestive tubule, int -intestine) (H&E, Scale bar = 100 µm). (p. 67). Figure 31A Ancistrum mytili ciliates (arrow) on the gills of a blue mussel (M. edulis). Note the loose association between the ciliates and the gill epithelium (compare with the Sphenophrya-like ciliates of eastern oyster (Figure 22B) and blue mussel (Figure I 32B). (H&E, Scale bar = 100 µm). (p. 69).

Figure 31B Specimen of A. mytili containing rickettsial-like hyperparasites (arrow). (H&E, I Scale bar = 20 µm). (p. 69).

Figure 32A Sphenophrya-like ciliate attached to the gills of a blue mussel (M. edulis). I Compare with Sphenophrya-like ciliates of eastern oyster (Figure 22B). (H&E, Scale bar = 100 µm). (p. 71).

I Figure 32B Sphenophrya-like ciliates inside an enlarged gill epithelial cell. Note distension of the cell nucleus (hn) as well as the cell itself. The ciliates can be seen with tentacle-like (arrows) filamentous extrusions into the host cell cytoplasm. (H&E, I Scale bar = 100 µm). (p. 71). Figure 33A "MPX" (mussel protozoan X) ciliates (arrows) inside the digestive tubule (dt) cells I of a blue mussel (1lf edulis). (H&E, Scale bar = 0.5 mm). (p. 73).

Figure 33B "MPX" ciliates lying within and between the tubule epithelial cells. Note dark I staining macronucleus (pn) compared to the granular nucleus of the tubule cells (en). Note also the vacuole-like space (vs) around many of the ciliates within I epithelial cells. (H&E, Scale bar = 50 µm). (p. 73). Figure 34A Ciliates (arrows) inside the intestine of a blue mussel (M. edulis). (H&E, Scale I bar = 100 µm). (p. 75). Figure 34B Intestinal ciliates showing large basophilic nuclei and polymorphic body shapes. Note the lack of attachment to the intestinal epithelium demonstrated by other internal ciliates (Figures 23A, B, and 33A, B). (H&E, Scale bar = 20 µm). (p. I 75). I Figure 35A Granuloma (g) associated with a pearl (p) encapsulating a digenean metacercaria (m) in the connective tissue of a blue mussel (M. edulis). (H&E, Scale bar = 100 I µm). (p. 77). I I 119

Figure 35B Pearl containing a digenean metacercaria in the connective tissue of a blue mussel (M. edulis) but with no associated granuloma. (H&E, Scale bar = 100 gm). (p. 77).

Figure 36A Copepod-like crustacean (c) in the cormective tissue of a blue mussel (M. edulis). Note the extensive granuloma (g) surrounding the crustacean, as well as throughout the digestive gland (arrows). (H&E, Scale bar = 0.5 mm) (p. 79).

Figure 36B Haemocyte response to a "stray" copepod within the tissues of a blue mussel (M. edulis). Note the granulocyte infiltration throughout the necrotic crustacean debris. (H&E, Scale bar = 100 gm). (p. 79).

CLAM PARASITES, PESTS AND DISEASES

Figure 37 Haematoxylin and eosin stained tissue-section through the tissues of a healthy quahaug (hard-shell clam) (Mercenaria mercenaria). Mantle (m), gonad (gd), gills (gl), digestive gland (dg), stomach (s), intestine (i), kidney (k) and foot (f). (Scale bar = 2 mm). (p. 82).

Figure 38A- Cross-section through quahaug (M. mercenaria) showing lesions caused by a thraustochytrid-like organism ("QPX" - Quahaug Parasite X) (arrows). (m.- mantle, dg - digestive gland, int - intestine). (H&E, Scale bar = 0.5 mm) (p. 83).

Figure 38B Multiplicative stages (ms) and daughter cells (dc) of "QPX" in the mantle tissue of quahaug (M. mercenaria). (H&E, Scale bar = 100 gm). (p. 83).

Figure 38C Colonies of Chytrid-like "Quahaug Parasite X" (QPX) cultured on Potato Dextrose Agar from necrotic quahaug (M. mercenaria) tissue. Thick-walled, cyst- like stages (arrows) are interspersed among smaller "daughter cell" stages. (Scale bar = 100 gm). (p. 84).

Figure 39A Cross-section through the digestive gland of a soft-shell clam (Mya arenaria) with haemic neoplasia. Note the characteristic enlarged and diffuse nuclei and corresponding reduction of cytoplasm in neoplastic cells (n) compared with normal haemocyte,s (h). Mitotic figures (m) are also present. Compare with haemic neoplasia of blue mussels (Figure 30B). (H&E, Scale bar = 100 iim). (p. 86).

Figure 39B Haemocyte suspension preparation from a healthy soft-shell clam (M. arenaria). Note the pseudopodial extensions of the normal haemocyte,s and small, granular nuclei. (Feulgen Picromethyl Blue (FPM), Scale bar = 20 pm). (p. 87).

Figure 39C Normal (n) and neoplastic (c) haemocytes in a haemolymph sample from a soft- shell clam (M. arenaria) with advanced haemic neoplasia. Note that most of the • neoplastic cells are rounded up and show no pseudopodial extensions of the cytoplasm. (FPM, Scale bar = 20 gm). (p. 87). 120 I Figure 40 Echinostome metacercaria (m) encysted within a pearl (p) in the gills of a quahaug (M. mercenaria). Note tiny spines (sp) around the anterior end of the body. I (H&E, Scale bar = 100 µm). (p. 89). Figure 41 Siphons of soft-shell clam (M. arenaria) infested by the marine snail Odostomia seminuda. The small white snails (< 3 mm long) are found clustered around the I opening of the inhalant siphon. (Scale bar = 5 mm). (p. 91). I SCALLOP PARASITES, PESTS AND DISEASES Figure 42A Haematoxylin and eosin stained tissue-section through the tissues of a healthy bay scallop (Argopecten irradians). Mantle (m), female gonad (fg), male gonad (mg), I gills (gl), digestive gland (dg), stomach (s), intestine (i) and kidney (k). (Scale bar = 2 mm). (p. 94).

I Figure 42B Haematoxylin and eosin stained tissue-sections of a healthy giant sea scallop (Placopecten magellanicus). Tissue sections of scallops > 75 mm wide will not fit on a single microscope slide (as shown in Figure 42A). Separate sections are I taken through the digestive gland (dg) and kidney (k), as well as the gills and gonad (Figure 42C). (Scale bar = 2 mm). (p. 95).

I Figure 42C Haematoxylin and eosin stained tissue-sections through the gills (gl) and gônad (gd) of a giant sea scallop (Placopecten magellanicus). (Scale bar = 2 mm). (p. I 95). Figure 43A Cross-section of bay scallop, Argopecten irradians, showing "swirl" lesions I induced by Perkinsus karlssoni. (H&E, Scale bar = 100 µm). (p. 96). Figure 43B High-power magnification of P. karlssoni within an early host haemocyte encapsulation. Note flattened haemocytes (h) around the parasites which are I characterised by the presence of vacuoles and dense inclusion bodies (ib). (H&E, Scale bar = 25 µm). (p. 96). I Figure 43C Lugol-positive spores of P. karlssoni, grown in thioglycollate medium from infected bay scallop (A. irradians) mantle tissue. (Scale bar = 100 µm). (p. 97). I Figure 44A Bay scallop, A. irradians, kidney infected by Pseudoklossia-like coccidian parasites (p). (rtl - renal tubule lumen; ke - kidney epithelium). (H&E, Scale bar = 100 µm). (p. 100).

I Figure 44B Pseudoklossia-like coccidians (p) between the epithelial cells of a bay scallop, A. irradians, stomach (es). (dt - digestive tubules, ct -connective tissue infiltrated by I haemocytes). (H&E, Scale bar = 100 µm). (p. 100). I I I I 121

I Figure 45 Echinostome metacercaria encysted within the kidney of a bay scallop, A. irradians. Note haemocyte encapsulation (ec) and spiny integument around the anterior end of the parasite. N.B. there is no evidence of pearl encapsulation (see I Figures 35B,C, 40). (H&E, Scale bar = 100 µm). (p. 102).

Figure 46 Unidentified digenean in the digestive gland of giant sea scallop, P. magellanicus, I infected by an unidentified digenean. Note unciliated, serrated integument (st), oral sucker (os) and haemocyte infiltration (h) into the connective tissue in front I of the parasite. (H&E, Scale bar = 100 µm). (p. 102). Figure 47A "Brown-spot" lesion (arrow) in the adductor muscle of giant sea scallop, I Placopecten magellanicus. (Scale bar = 1 mm). (p. 1(4). Figure 47B Cross-section through giant sea scallop, P. magellanicus, muscle fibres (mf), affected by "brown spot". The abscess lesions are filled with necrotic haemocytes I (nh) and muscle fibres (nm). (H&E, Scale bar = 100 µm). (p. 104). Figure 48A Cross-section of giant sea scallop, P. magellanicus, gills with Trichodina-like I ciliates (arrows). (H&E, Scale bar = 100 µm). (p. 106). Figure 48B Trichodinid-like ciliate on the gills of giant sea scallop, P. magellanicus. (H&E, I Scale bar = 20 µm). (p. 106). Figure 49A Cross-section of giant sea scallop, P. magellanicus, gills and a surface I turbellarian. (H&E, Scale bar = 0.5 mm). (p. 108). Figure 49B Turbellarian from the gills of giant sea scallop, P. magellanicus magnified toshow 1 ciliated epidermis. (H&E, Scale bar = 100 µm). (p. 108). Figure 50 Cross-section through the intestine (int) of giant sea scallop, P. magellanicus, containing a turbellarian (tb). Note ciliated epidermis (ce) and pharynx (ph). I (H&E, Scale bar = 100 µm). (p. 110). I I I I I I I I I I

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