Histological Techniques for Marine Bivalve 

Molluscs: Update 

 

 

NOAA Technical Memorandum NOS NCCOS 27 
Mention of trade names or commercial products does not constitute 

endorsement or recommendation for their use by the United States 

government. 

Cover photograph of trematode sporocysts completely occupying the mantle of a mytilid 

mussel, Mytilus edulis. 

Citation for this Report 

Kim, Y., K.A. Ashton-Alcox, and E.N. Powell. 2006. Histological Techniques for Marine 

Bivalve Molluscs: Update. Silver Spring, MD. NOAA Technical Memorandum NOS 

NCCOS 27. 76 pp.
Histological Techniques for Marine Bivalve 

Molluscs: Update 

 

Kim, Y., K. A. Ashton-Alcox, and E. N. Powell 

Center for Coastal Monitoring and Assessment (CCMA) 

NOAA/NOS/NCCOS 

1305 East-West Highway 

Silver Spring, Maryland 20910 

NOAA Technical Memorandum NOS NCCOS 27 

May 2006 

United States Department of National Oceanic and National Ocean Service 

Commerce Atmospheric Administration 

Carlos M. Gutierrez Conrad C. Lautenbacher, Jr. Jack Dunnigan 

Secretary Administrator Assistant Administrator 

3

TABLE OF CONTENTS 

TABLE OF CONTENTS .................................................................................................. i

LIST OF TABLES ...........................................................................................................iii

LIST OF FIGURES ......................................................................................................... iv

CHAPTER 1. GONADAL ANALYSIS .......................................................................... 1

ABSTRACT.................................................................................................................... 1 

1.0 INTRODUCTION .............................................................................................. 1 

2.0 EQUIPMENT, REAGENTS, SOLUTIONS, AND SAMPLE PREPARATION

 ...............................................................................................2 

3.0 SAMPLE COLLECTION AND FIXATION ..................................................... 5 

4.0 SLIDE PREPARATION..................................................................................... 7 

5.0 ANALYSIS......................................................................................................... 9 

6.0 CONCLUSIONS............................................................................................... 14 

7.0 REFERENCES ................................................................................................. 15 

CHAPTER 2. HISTOPATHOLOGY ANALYSIS..................................................... 19

ABSTRACT.................................................................................................................. 19 

1.0 INTRODUCTION ............................................................................................ 19 

2.0 EQUIPMENT, REAGENTS, AND SOLUTIONS........................................... 20 

3.0 ANALYSIS....................................................................................................... 20 

4.0 CONCLUSIONS............................................................................................... 43 

5.0 REFERENCES ................................................................................................. 43 

CHAPTER 3. PERKINSUS MARINUS ASSAY ......................................................... 53

ABSTRACT.................................................................................................................. 53 

1.0 INTRODUCTION ............................................................................................ 53 

2.0 EQUIPMENT AND SUPPLIES....................................................................... 54 

i
3.0 TISSUE COLLECTION................................................................................... 56 

4.0 TISSUE ANALYSIS ........................................................................................ 56 

5.0 DEFINITIONS AND CALCULATIONS ........................................................ 60 

6.0 CONCLUSIONS............................................................................................... 61 

7.0 REFERENCES ................................................................................................. 61 

ii
LIST OF TABLES 

Table 1-1. Tissue embedding sequence. ............................................................................. 8 

Table 1-2. Tissue staining sequence. ................................................................................ 11 

Table 1-3. Oyster development stages adapted from Ford and Figueras (1988) by Powell 

et al. (1993)............................................................................................................... 12 

Table 1-4. Mytilid and dreissenid development stages adapted from Seed (1975, 1976) by 

Hillman (1993).......................................................................................................... 13 

Table 1-5. Semi-quantitative scale for abnormal gonadal development in mytilid mussels.

................................................................................................................................... 14 

Table 2-1. List of quantitative and semi-quantitative categories for each bivalve taxon.. 22 

Table 2-2. Semi-quantitative scale for Haplosporidium nelsoni infection modified from 

Ford (1985, 1986), and Ford and Figueras (1988).................................................... 35 

Table 2-3. Composite rating matrix for Haplosporidium nelsoni infection. .................... 36 

Table 2-4. Semi-quantitative scale for trematode sporocyst infection. ............................ 38 

Table 2-5. Semi-quantitative scale for digestive gland atrophy. ...................................... 40 

Table 3-1. Oyster condition rating key (from Quick and Mackin, 1971)......................... 57 

Table 3-2. Semi-quantitative scale of infection intensity for Perkinsus marinus [adapted 

from Mackin (1962) by Craig et al. (1989)]. ............................................................ 59 

iii
LIST OF FIGURES 

Figure 1-1. Mytilus edulis follicle with abnormal gametic tissue infiltrated with 

hemocytes. ................................................................................................................ 15 

Figure 2-1. Prokaryotic inclusions present in digestive tract epithelium of an oyster, 

Crassostrea virginica................................................................................................ 24 

Figure 2-2. Cyst-like encapsulations of prokaryotic microorganism in gills of a mytilid 

mussel, Mytilus edulis............................................................................................... 24 

Figure 2-3. Nematopsis sp. spores in connective tissue between digestive tubules of an 

oyster, C. virginica. Arrows indicate examples........................................................ 25 

Figure 2-4. Numerous Nematopsis sp. spores in the gills of an oyster, C. virginica. 

Arrows indicate examples. ....................................................................................... 25 

Figure 2-5. Gregarine-like spores near the tip of the mantle of a mytilid mussel, M. 

californianus. Arrows indicate examples.. ............................................................... 26 

Figure 2-6. Ciliates between the gill filaments of a mytilid mussel, M. edulis. Arrows 

indicate examples...................................................................................................... 27 

Figure 2-7. Ciliates in the lumen and attached to the epithelium of the intestine of an 

oyster, C. virginica. Arrows indicate examples........................................................ 28 

Figure 2-8. Ciliated xenomas on the gill surface of an oyster, C. virginica. Arrows 

indicate examples...................................................................................................... 28 

Figure 2-9. Encapsulated larval cestodes in the vesicular connective tissue surrounding 

the stomach of an oyster, C. virginica.. .................................................................... 29 

Figure 2-10. A larval cestode in the gill connective tissue of an oyster, C. virginica...... 30 

Figure 2-11. Cross-sections of metacercariae encysted in the visceral connective tissue of 

a mytilid mussel, M. edulis.. ..................................................................................... 31 

Figure 2-12. Sections of unidentified nematode larvae in the digestive gland connective 

tissue of an oyster, C. virginica. Arrows indicate examples..................................... 31 

Figure 2-13. Neoplastic cells infiltrating the visceral connective tissue of a mytilid 

mussel, M. edulis....................................................................................................... 33 

Figure 2-14. Hemocytic infiltration near the gill base of an oyster, C. virginica. Arrows 

indicate examples...................................................................................................... 34 

Figure 2-15. Granulocytomas in the digestive gland of a mytilid mussel, M. edulis. 

Arrows indicate examples......................................................................................... 34 

iv
Figure 2-16. Numerous multinucleated plasmodia of Haplosporidium nelsoni in the gills 

of an oyster, C. virginica. Arrows point to example parasites.................................. 36 

Figure 2-17. This trematode sporocyst infection in Mytilus edulis scored a 1 according to 

Table 2-4.. ................................................................................................................. 38 

Figure 2-18. This trematode sporocyst infection in Mytilus edulis scored a 2 according to 

Table 2-4. No gametic tissue is present. ................................................................... 39 

Figure 2-19. Crassostrea virginica normal digestive tubule, scored a 0 according to Table 

2-5. ............................................................................................................................ 40 

Figure 2-20. Digestive gland atrophy in Crassostrea virginica scored a 2 according to 

Table 2-5. .................................................................................................................. 41 

Figure 2-21. Digestive gland atrophy in Crassostrea virginica scored a 4 according to 

Table 2-5. .................................................................................................................. 41 

Figure 2-22. Unusual digestive tubules of a mytilid mussel, M. edulis............................ 43 

Figure 3-1. Enlarged and stained Perkinsus marinus hypnospores. Hypnospores appear as 

black dots in the photographs. .................................................................................. 58 

v

CHAPTER 1. GONADAL ANALYSIS 

Y. Kim, K. A. Ashton-Alcox, and E. N. Powell 

Haskin Shellfish Research Laboratory 

Rutgers University 

Port Norris, NJ 08349 

ABSTRACT 

This chapter describes the procedures for determining the reproductive stage of oysters, 

mytilid mussels, and dreissenid mussels collected for NOAA’s National Status and 

Trends Mussel Watch Project. Analyses are conducted on paraffin-embedded tissues 

sectioned at a 5-μm thickness and stained using a pentachrome staining procedure. Each 

slide is examined microscopically to determine the animal’s sex and stage of gonadal 

development. A semi-quantitative ranking is assigned. 

1.0 INTRODUCTION 

Assessment of the physiological state of bivalve populations requires an analysis of the 

state of gonadal development. Determination of reproductive stage is included as part of 

the Mussel Watch Project to give an indication of the amount of gametic material in 

bivalve tissues at the time of chemical analysis. Certain contaminants are preferentially 

concentrated in gonadal tissue (Ellis et al., 1993; Lee, 1993; Abbe et al., 1994). Others 

are concentrated in non-gonadal tissue (Cunningham and Tripp 1975; Mo and Neilson, 

1993). Because gametic material can account for 20% to 50% of body weight in target 

species of oysters and mussels (Sprung, 1991; Choi et al., 1993), the relative proportion 

of gonadal to somatic tissue and the timing of spawns (an important depuration route for 

some contaminants) can significantly impact the body burden of contaminants. 

This description updates the methods presented in Ellis et al. (1998b) and Powell et al. 

(1993). The original intent of the determination of reproductive stage was to assure that 

sampling was conducted at the same stage of the reproductive cycle so that analyses of 

the lipophilic organic contaminants and the trace element contaminants were not 

influenced by reproductive state. Unfortunately, the time required for sampling and the 

wide latitudinal range encompassed by the sites did not permit consistent recovery of 

individuals in similar stages of reproductive development at all sites. For example, 

typically oysters are undifferentiated in the winter. Gonads begin to develop in early 

spring and spawning occurs late spring through early fall. Most Gulf Coast oysters spawn 

at least twice during this time period. Single spawns tend to occur in the shorter summers 

of the mid-Atlantic region (e.g., Dittman et al., 2001). The timing of the last spawn varies 

with latitude and with yearly variations in climate (e.g., Wilson et al., 1990, 1992). 

Southeast Atlantic and Southern Gulf sites, for example, routinely yield oysters in 

reproductive development or that are ready to spawn in mid-winter during the period 

when Mussel Watch sampling occurs. Mid-Atlantic sites are typically characterized by 

individuals in an undifferentiated state and thus contain significantly less lipid-rich 

1
gametic tissue than the southern animals. Mytilid mussels and dreissenid mussels have 

the same assortment of problems relating to latitude and interannual changes in climate 

(Newell, 1989; Seed and Suchanek, 1992; Borcherding, 1991). In addition, dreissenid 

mussels are typically collected during late August-September whereas the remaining 

Mussel Watch species are collected during winter. Thus, analysis of reproductive stage 

has proved important in identifying differences in tissue composition that might affect 

comparisons of contaminant data among sites and among years. Wilson et al. (1990, 

1992) and Kim et al. (1999, 2001) discuss the influence of climate on reproductive stage 

and contaminant body burden in the Mussel Watch Project in more detail. 

Oyster gonadal tissue is distributed around the body mass (Morales-Alamo and Mann, 

1989). Gonads of dreissenid mussels also develop within the body, around the periphery 

of the viscera. In contrast to oysters and dreissenid mussels, gonadal follicles develop 

primarily within the mantle of mytilid mussels. Nevertheless, in none of these cases can 

the gonad be easily excised and weighed. Consequently, virtually all assays of 

reproductive stage use histological methods to recognize the changes in the germinal 

epithelium and germinal products that identify stages in gonadal development. The 

histological approach uses a semiquantitative numerical assignment to rank reproductive 

stage. Quantitative measures, such as egg protein content (Choi and Powell, 1993, Choi 

et al., 1993, 1994), remain expensive and time consuming, and do not permit a 

concomitant histopathological analysis. Therefore, a histological examination is still the 

single method of choice when only one method can be used. 

For oysters, a dorsal-ventral slice of tissue is taken and fixed in Davidson’s fixative. 

Dreissenid and mytilid mussels are preserved whole, and a dorsal-ventral slice is taken 

after fixation. Tissue slices are embedded in paraffin, sectioned, and stained using a 

pentachrome staining protocol. Stained sections are examined under a compound 

microscope, and sex and the state of gonadal development determined. Fixation follows 

the method described in Preece (1972). The staining procedure is an adaptation of 

Masson’s (1928) trichrome procedure (Ellis et al., 1998b). Reproductive stage in oysters 

is determined using a semiquantitative scale adapted from Ford and Figueras (1988). The 

scale developed by Seed (1975, 1976) for determining gonad index in mussels was 

adopted for mytilid mussels and dreissenid mussels. 

2.0 EQUIPMENT, REAGENTS, SOLUTIONS, AND SAMPLE 

PREPARATION 

2.1. EQUIPMENT 

Cover slips - various sizes. Fisher Scientific, Pittsburgh, PA. 

Disposable cassettes - HistoPrep tissue capsules, 15-182-218. Fisher Scientific, 

Pittsburgh, PA. 

Disposable microtome blades, 12-634-11. Fisher Scientific, Pittsburgh, PA. 

Drying oven, 13-254-1. Fisher Scientific, Pittsburgh, PA. 

2
Embedding rings - HistoPrep embedding rings, 12-652-1OB. Fisher Scientific, 

Pittsburgh, PA. 

Frosted microslides, 12-552. Fisher Scientific, Pittsburgh, PA. 

Microscope, compound, Zeiss, 12-070-20. Fisher Scientific, Pittsburgh, PA. 

Microtome, HM330. Microm GmbH. D-6900 Heidelberg, Germany. 

Slide staining set, Tissue-Tek II. Miles Laboratories, Inc. Westmont, IL. 

Slide staining holder, Tissue-Tek, 4466. Miles Laboratories, Inc. Westmont, IL. 

Stainless steel molds, various sizes. Fisher Scientific, Pittsburgh, PA. 

Tissue embedding system, Histocentre 2, 6400012. Shandon, Inc., Pittsburgh, PA. 

Tissue processor, Citadel 2000, 69800006. Shandon, Inc., Pittsburgh, PA. 

Vacuum infiltrator, Tissue-Tek II, 4613. Miles Laboratories, Inc. Westmont, IL. 

Water bath - Fisher Tissue PrepModel 134 Flotation Bath, 15-464. Fisher Scientific, 

Pittsburgh, PA. 

2.2. REAGENTS 

Acetone HistoPrep (CH3COCH3), [67-64-1], HC300-1GAL. Fisher Scientific, Pittsburgh, 

PA. 

Acid fuchsin, certified stain (C20H17N3O9S3Ca) [136132-76-8], A3908. Sigma Chemical 

Co.,St. Louis, MO. 

Ammonium hydroxide (NH4OH) [1336-21-6], A6899. Sigma Chemical Co., St. Louis, 

MO. 

Aniline blue powder, certified stain [28631-66-5], A967-25. Fisher Scientific, Pittsburgh, 

PA. 

Chromotrope powder 2R, (C16H10N2Na2O8S2) [4197-07-3], C3143. Sigma Chemical 

Co., St. Louis, MO. 

Ethanol (C2H5OH) [64-17-5], R8382. Sigma Chemical Co., St. Louis, MO. 

Fast green FCF, certified stain (C37H34N2O10S3Na2) [2353-45-9], F7252. Sigma 

Chemical Co., St. Louis, MO. 

3
Ferric ammonium sulfate (Fe NH4(SO4)2 . 12H2O) [7783-83-7], F1018. Sigma Chemical 

Co., St. Louis, MO. 

Formaldehyde, 37% solution (CH2O) [50-00-0], F1635. Sigma Chemical Co., St. Louis, 

MO. 

Glacial acetic acid (C2H4O2) [64-19-7], A0808. Sigma Chemical Co., St. Louis, MO. 

Glycerin (C3H8O3) [56-81-5], G7893. Sigma Chemical Co., St. Louis, MO. 

Hematoxylin powder, certified stain (C16H14O6) [517-28-2], H3136. Sigma Chemical 

Co., St. Louis, MO. 

Orange G powder, certified stain (C16H10N2O7S2Na2) [1936-15-8], O7252. Sigma 

Chemical Co., St. Louis, MO. 

Paraffin - Paraplast tissue embedding media (melting pt. 56 °C), 12-646-111, Fisher 

Scientific, Pittsburgh, PA. 

Permount mounting media, SP15-500. Fisher Scientific, Pittsburgh, PA. 

Phosphomolybdic acid (20MoO3 . 2H3PO4 . 48H2O) [51429-74-4], P7390. Sigma 

Chemical Co., St. Louis, MO. 

Phosphotungstic acid (12WO3 . H3PO4 . H2O) [12501-23-4], P4006. Sigma Chemical 

Co., St. Louis, MO. 

Sodium chloride (NaCl) [7647-14-5], S9625. Sigma Chemical Co., St. Louis, MO. 

Sulfuric acid (H2SO4) [7664-93-9], S1526. Sigma Chemical Co., St. Louis, MO. 

Tissue Clear III, SH3-4. Fisher Scientific, Pittsburgh, PA. 

Tissue Dry, SH5-20. Fisher Scientific, Pittsburgh, PA. 

Xyleness, histological grade (C6H4(CH3)2), X3S-4. Fisher Scientific, Pittsburgh, PA. 

2.3. SOLUTIONS 

Ferric alum mordant: 25 g ferric ammonium sulfate dissolved in 500 mL distilled water. 

Basic ethanol: 26 mL ammonium hydroxide in a solution of 3370 mL 95% ethanol and 

630 mL distilled water. 

4
Phosphomolybdic acid solution: 5 g phosphomolybdic acid crystals dissolved in 495 mL 

distilled water. 

1% acetic acid: 20 mL glacial acetic acid in 1980 mL distilled water. 

1% acid acetone: 20 mL glacial acetic acid in 1980 mL acetone. 

Groat/Weigert hematoxylin working solution: 245 mL distilled water, 5 mL sulfuric acid, 

5 g ferric ammonium sulfate, 245 mL 95% ethanol, and 2.5 g hematoxylin powder. 

Acid fuchsin working solution: 1.5 g acid fuchsin powder dissolved in 495 mL distilled 

water, to which is added 5 mL glacial acetic acid. 

Phosphotungstic acid solution: 10 g phosphotungstic acid crystals dissolved in 490 mL 

distilled water. 

Orange G/Chromotrope: 4 g orange G powder and 1 g chromotrope powder dissolved in 

495 mL distilled water to which was added 5 mL glacial acetic acid. 

Fast green/Aniline blue working solution: 5 g fast green FCF, 4 g aniline blue powder 

dissolved in 495 mL distilled water to which was added 5 mL glacial acetic acid. 

Davidson's fixative solution: 1 part glycerin, 1 part glacial acetic acid, 2 parts 37% 

formaldehyde, 3 parts 95% ethanol, and 3 parts isotonic sodium chloride (usually 20 - 

30‰). 

3.0 SAMPLE COLLECTION AND FIXATION 

3.1. SAMPLING 

From 1986 to 1994, the same oysters were used for organic contaminant analysis and 

gonadal analysis at all Gulf coast sites (e.g., Powell et al., 1993). The use of the same 

animals for gonadal analysis and for analysis of contaminant body burden potentially 

biases the latter analyses because digestive gland tissue and gonadal tissue, that 

contribute disproportionately to the tissue taken for histological analysis, may contain a 

higher than average body burden of certain contaminants. Sericano et al. (1993) showed 

that this source of error resulted in an underestimation of true body burden no greater 

than 10% when a 5-mm slice was removed from the large oysters normally sampled in 

the Mussel Watch Project. This potential error would be much larger for the smaller 

mytilids and dreissenids. To avoid this error, separate samples have always been obtained 

for gonadal analysis of mussels and East Coast oysters. Beginning in 1995, the same 

protocol was adopted for Gulf Coast oysters. The number of animals analyzed per site 

has also changed. Through 1994, 15 animals were analyzed per site. Thereafter, five were 

analyzed per site. Thus, the present sampling method requires the assumption that 

individuals collected from a common collection area will have experienced similar 

5
chemical loading and that the five animals analyzed for gonadal analysis are 

representative of the animals pooled for contaminant analysis. 

3.2. TISSUE PREPARATION 

3.2.1. OYSTER TISSUE PREPARATION

Five of the 12 oysters processed for Dermo analysis (Ashton-Alcox et al., this volume) 

are chosen randomly for gonadal analysis. A 5-mm thick cross-section of tissue is 

removed from the oyster using a scalpel or scissors. The determination of reproductive 

stage is based on a histological evaluation of the maturation stage of oyster gonads 

located within/around the visceral mass. The tissue section is obtained such that the 

dorsal-ventral aspect passes through the digestive gland and gill tissue just posterior to 

the palps (Figure 1 in Ellis et al., 1998). This aspect provides example sections of most 

oyster tissues for histopathological analysis (Kim et al., this volume), while also 

providing a representative cross-section of oyster gonad. Each section is immediately 

placed in a tissue cassette and the cassette placed in a jar filled with Davidson’s fixative 

for 48 hr. After 48 hr, the fixative is decanted, 70% ethanol added and the tissues are 

allowed to stand until processing. 

3.2.2. MYTILID MUSSEL TISSUE PREPARATION

Determination of reproductive stage for mussels is based on a histological evaluation of 

the maturation stages of mussel gonads, most of which are located in the mantle (Newell, 

1989). Five animals are analyzed but a few more are preserved for reasons discussed 

subsequently. The tip of a sharp knife is carefully inserted between the shells at the 

ventral lip and run dorsally between the shells until the posterior adductor muscle is cut 

so that the shells remain in an open position. Care is taken to cut no further than the 

adductor muscle to avoid cutting into the digestive gland immediately below the adductor 

muscle. Shucking of fresh mussels usually results in severe damage to the mantle tissue 

lying next to the shell. Therefore the mussels are placed whole in a wide-mouth jar filled 

with Davidson’s fixative after the adductor muscle has been cut. Because the entire 

animal is being preserved, the specimens are left in fixative for at least a week to ensure 

preservation of all tissues. After this time, the fixative is decanted and 70% ethanol is 

added for storage until processing. 

Once preserved, the tissue hardens and becomes easier to detach from the shell. To excise 

the preserved mussel meat from its shell, a knife is carefully run between the mantle and 

the lip of each valve, detaching the mantle from the shell. At this time, byssal threads are 

completely removed from the byssal gland to avoid later difficulties in tissue sectioning. 

Five specimens are chosen from each site and their anterior-posterior lengths are 

measured using a ruler. A 5-mm thick cross-section is then removed using a scalpel. The 

cross-section is obtained such that the dorsal-ventral aspect passes through the digestive 

gland and gills at an angle across the body and such that ventral edge of the cross-section 

is slightly towards the posterior-ventral margin. Each cross-section is placed in a tissue 

cassette and processed immediately after dissection. If the mantle tissue is damaged 

during the shucking procedure, the specimen is replaced by one of the additional 

6
specimens preserved from the same site because the wound could result in the loss of 

gametic material and lead to an erroneous evaluation. 

3.2.3. DREISSENID MUSSEL TISSUE PREPARATION

Most of the gonad of a dreissenid mussel is concentrated within the visceral mass 

(Borcherding, 1990). Due to their small size however, dissection of living tissue without 

destroying the gonads is difficult. Therefore, dreissenid mussels collected from each site 

are preserved whole in Davidson’s fixative, without cutting the adductor muscle. They 

are left in fixative for one week to allow adequate time for tissue fixation. After this time, 

20 to 30 mL of acetic acid is added to enhance decalcification of the shell. The shell is 

properly decalcified when it is no longer hard. 

After decalcification, the Davidson’s fixative is replaced with 70% ethanol according to 

the procedure followed for mytilid mussels and stored for later embedding. Prior to 

embedding, byssal threads are cut away from the byssal gland to minimize difficulty in 

sectioning the tissue. A 5-mm thick cross-section is taken from five individuals as 

described for mytilid mussels. Each section is placed in a tissue cassette and processed 

for embedding immediately thereafter. 

4.0 SLIDE PREPARATION 

4.1. TISSUE EMBEDDING 

Individual tissue samples are prepared for embedding in paraffin using an established 

dehydration protocol (Table 1-1). The solutions used for dehydration, clearing, and 

infiltration are changed frequently to maintain solution purity. The tissue embedding 

sequence uses an automated tissue processor that processes tissue in plastic cassettes 

through the dehydration-clearing series and into paraffin. Embedding can also be done 

manually by moving the tissues through the sequence. The paraffin is melted in an 

embedding center with temperature set at 60°C. Newly melted paraffin should always be 

used in the final infiltration and embedding steps. 

After the tissues are infiltrated with paraffin (Table 1-1), they are transferred to a vacuum 

infiltrator set at 60°C and placed under a vacuum for 30 min. Tissues are transferred to a 

holding tank of melted paraffin and removed singly to stainless steel molds. The tissues 

are oriented with the cross-sectional face down for sectioning, and a plastic mold 

embedding ring is placed on top. The ring is filled with paraffin and the mold moved to a 

cold plate of the embedding system. As the tissue/paraffin cools and hardens, the paraffin 

shrinks. Care must be taken to use sufficient paraffin to cover the tissue after this 

shrinkage. The mold is left on the cold plate until the tissue-paraffin block is removed. 

The block is then placed in a freezer until sectioning. 

7
4.2. TISSUE SECTIONING 

The paraffin blocks are first cut at 20 μm to expose an entire tissue cross-section and then 

sliced at 5 μm using a microtome. Tissue sections may be cut singly or into contiguous 

sections. The sections are placed on the surface of a water bath maintained at 45-50°C 

and allowed to expand. Once the sections expand to full size, a microscope slide is held at 

an angle and slid under one or more of the tissue sections. The sections are then lifted out 

of the water and onto the slide. The sections are positioned on the slide in the orientation 

in which they will be stained and read. The slide is allowed to air dry until it can be 

placed in a slide rack. The slide rack is placed in a drying oven at 40°C. After drying 

overnight or longer, the slides are ready to stain. 

Table 1-1. Tissue embedding sequence*. 

________________________________________________________________________ 

Dehydration Clearing 

 Tissue Dry 60 min Tissue Clear 60 min 

 Tissue Dry 120 min Tissue Clear 120 min 

 Tissue Dry 120 min Tissue Clear 120 min 

 Tissue Dry 120 min 

 Tissue Dry 120 min Infiltration 

 Tissue Dry 120 min 

 Tissue Dry 60 min paraffin 120 min 

 paraffin 120 min 

paraffin (in vacuum infiltrator) 30min 

 

* In cases where the sequential solutions are the same, each transfer is a transfer to a 

fresh solution. 

4.3. TISSUE STAINING 

Sections are deparaffinized and hydrated using a xylenes-ethanol series (Table 1-2). 

Following hydration, slides are stained in a pentachrome series, dehydrated in a series of 

acetic acid dips followed by acetone, cleared in xyleness and mounted in Permount 

(Table 1-2). The pentachrome staining procedure is an adaptation of the trichrome stain 

of Masson (1928) as modified by Gurr (1956) (Ellis et al., 1998b). The modifications 

include the addition of aniline blue to the fast green working solution, substitution of 

chromotrope 2R/orange G for Ponceau de Xylidene, and the addition of phosphotungstic 

acid prior to the orange G/chromotrope stain; the procedure is now a pentachrome 

technique. The addition of these stains yields better differentiation of tissue types and 

mucins. Times required for each step are flexible in both the staining procedures 

discussed here and in the previous embedding protocol. Different tissue types may 

require different times. All solutions, especially the xyleness and ethanol ones, should be 

changed frequently. Slides should not be allowed to dry during transfers. Solutions to 

8
common embedding, sectioning, and staining problems are discussed in Preece (1972) 

and most other manuals of histological technique. 

5.0 ANALYSIS 

Each slide is examined microscopically to determine sex and stage of gonadal 

development. A histopathological examination can also be made at this time (Kim et al., 

this volume). Careful examination of early developmental stages is needed to positively 

distinguish males or females in early stages of development from individuals as yet 

undifferentiated. Occasional hermaphrodites will also be found (all target species 

normally have separate sexes). The stage in the gametogenic cycle is assigned based on 

the maturity of the follicles and gametes and a numerical value is assigned as described in 

Tables 1-3 and 1-4. 

Cases of renewed gonadal development following spawning are common in oysters 

(stage 7), particularly along the Gulf of Mexico coast (Supan and Wilson, 2001). These 

animals typically have a few remaining large, mature ova and many developing ova that 

would normally be found in stages 3 or 4. Accordingly, for oysters, further data reduction 

can better be achieved by comparing the number of individuals with substantial gonadal 

development with those having little gonadal volume using an egg/eggless ratio, 

calculated as: 

the number of individuals at stages 3, 4, 5, 6 and 7 

Oyster egg/eggless ratio = the number of individuals at stages 1, 2 and 8 

For mytilids and dreissenids, the analogous ratio is calculated as: 

the number of individuals at stages 3, 4 and 5 

Mussel egg/eggless ratio = the number of individuals at stages 0, 1 and 2 

Abnormal gonadal development is commonly observed in mytilid mussels. This is often 

characterized by unusual development of gametes at the base of the follicles. The cells 

resemble those of a germinoma (Peters et al. 1994) and are differentiated from normal 

cells by being either enlarged or by appearing to have an enlarged nucleus. In other cases, 

underdeveloped, small gonadal follicles are observed. These occupy a smaller portion of 

the mantle tissue. Follicles may be filled with cellular debris (Figure 1-1; see also Figure 

10 in Ellis et al., 1998a). Sometimes cells adhere to each other, forming accumulations 

and empty spaces among developing cells. Occasionally, fibrosis occurs, with 

proliferation of fibroblasts inside the follicles and in the interfollicular connective tissue. 

Abnormal gonadal development is often associated with degeneration of Leydig tissue 

around the follicles and hemocytic infiltration into the surrounding tissues (Figure 11 in 

Ellis et al., 1998). The approach used to score instances of abnormal gonadal 

9
development uses a scale that rates the spatial coverage of the condition (e.g., fraction of 

follicles affected), but not the degree of effect in each follicle (Table 1-5). Normally, the 

entire follicle is completely affected or unaffected. 

10
Table 1-2. Tissue staining sequence. 

Deparaffinization 

xylenes 5 min 

xylenes 5 min 

xylenes 5 min 

100% ethanol 3 min 

100% ethanol 2 min 

Hydration 

95% ethanol 2 min 

10% ethanol 2 min 

distilled water 2 min 

Staining series 

Ferric alum mordant 10 min 

Running tap water quick dip 

Groat/Weigert Hematoxylin* 30-45 min 

Running tap water 5 min 

Acid Fuchsin stain 1.5 min 

Running tap water** 5 min 

Phosphotungstic acid 2 min 

Orange G/Chromotrope stain 1.5 min 

Running tap water** 5 min 

Phosphomolybdic acid 2 min 

Fast Green/Aniline Blue stain 3 min 

Dehydration 

1% acetic acid 20-25 dips 

1% acetic acid 20-25 dips 

1% acetic acid 20-25 dips 

1% acid acetone 20-25 dips 

1% acid acetone 20-25 dips 

1% acid acetone 20-25 dips 

 

Clearing 

xylenes 5 min 

xylenes 5 min 

 

Mounting 

 Mounting in Permount 24 hr to dry 

* A basic ethanol dip can be used to blue the hematoxylin, if necessary. 

** At these steps, no stain should remain between the slides and holding grooves in the slide 

rack. 

11
Table 1-3. Oyster development stages adapted from Ford and Figueras (1988) by Powell 

et al. (1993). 

Developmental Stage Value Description 

Sexually undifferentiated 1 Little or no gonadal tissue visible 

Early development 2 Follicles beginning to expand 

Mid development 3 Follicles expanded and beginning to coalesce; no 

mature gametes present 

Late development 4 Follicles greatly expanded, and coalesced, but 

considerable connective tissue remaining; some 

mature gametes present 

Fully developed 5 Most gametes mature; little connective tissue 

remaining 

Spawning 6 Gametes visible in gonoducts 

Spawned 7 Reduced number of gametes; some mature gametes 

still remaining; evidence of renewed reproductive 

activity 

Spawned 8 Few or no gametes visible; gonadal tissue 

atrophying 

12
Table 1-4. Mytilid and dreissenid development stages adapted from Seed (1975, 1976) by 

Hillman (1993). 

Reproductive stage Description 

Resting/spent gonad 

Stage 0 Inactive or undifferentiated 

Developing gonad 

 Stage 1 Gametogenesis has begun; no ripe gametes visible 

 Stage 2 Ripe gametes present; gonad developed to about one-third of its 

final size 

 Stage 3 Gonad increased in mass to about half the fully ripe condition; 

each follicle contains, in area, about equal proportions of ripe and 

developing gametes 

 Stage 4 Gametogenesis still progressing, follicles contain mainly ripe 

gametes 

Ripe gonad 

Stage 5 Gonad fully ripe, early stages of gametogenesis rare; follicles 

distended with ripe gametes; ova compacted into polygonal 

configurations; sperm with visible tails 

Spawning gonad 

 Stage 4 Active emission has begun; sperm density reduced; ova rounded 

off as pressure within follicles is reduced 

 Stage 3 Gonad about half empty 

 Stage 2 Gonadal area reduced; follicles about one-third full of ripe gametes 

 Stage 1 Only residual gametes remain; some may be undergoing cytolysis 

13
Table 1-5. Semi-quantitative scale for abnormal gonadal development in mytilid mussels. 

Score Description 

0 Normal gonad 

1 Less than half the follicles are affected 

2 About half the follicles are affected 

3 More than half the follicles are affected 

4 All follicles affected 

6.0 CONCLUSIONS 

The procedures described provide a semiquantitative ranking of reproductive stage 

but no quantification of the amount of gametic tissue present. The strengths of this 

approach are that it provides an assessment of sexual stage in the gametogenic cycle 

and allows for a concomitant histopathological analysis, with a single sample 

preparation protocol. The procedure cannot be performed on pooled samples. Thus, a 

direct correspondence between, for example, hydrocarbon body burden and stage in 

the gametogenic cycle may be difficult, because subsampling of individual animals 

will result in a certain degree of bias in the measurement of contaminant body burden, 

normally around 10% in adult oysters (Sericano et al., 1993) and more for smaller 

individuals and species. This bias, therefore, will be size, contaminant, and time-ofyear dependent.

If a quantitative gonadal/somatic index is desired, the technique of Choi and Powell 

(1993) should be used. The latter technique measures the concentration of egg protein 

present; however, it is not compatible with a concomitant histopathological analysis 

in that the standard histological preparation for assessing reproductive stage is not 

used in the quantitative analysis and tissue subsampling for histology cannot be done 

on the same individuals to be analyzed quantitatively for gonadal-somatic index. Choi 

et al. (1993) have further discussed the relative value of various approaches to 

gonadal evaluation. Overall, these authors found that the same general trends could be 

identified using either the semiquantitative or quantitative technique in most species, 

because normally an increase in gonadal volume occurs more or less simultaneously 

with advancement in reproductive stage and because gamete maturation occurs more 

or less evenly throughout the entire gonad. 

14
Figure 1-1. Mytilus edulis follicle with abnormal gametic tissue infiltrated with 

hemocytes. 

7.0 REFERENCES 

Abbe, G. R., J. G. Sanders and G. F. Riedel (1994) Silver uptake by the oyster 

(Crassostrea virginica): effect of organisms size and storage sites. Estuarine Coastal 

Shelf Sci., 39:249-60. 

Borcherding, J. (1990) Die Reproduktionsleistungen der Wandermuschel Dreissena 

polymorpha. Ph.D. Dissertation, Universität zu Köln, Germany. 168 pp. 

Borcherding, J. (1991) The annual reproductive cycle of the freshwater mussel Dreissena 

polymorpha Pallas in lakes. Oecologia (Berl.), 87:208-18. 

Choi, K-S. and E. N. Powell (1993) Development of and immunological probe for 

quantification of oyster gonad. In: Sampling and analytical methods of the National 

Status and Trends Program National Benthic Surveillance and Mussel Watch Projects 

1984-1992. Vol. II Comprehensive descriptions of complementary measurements. 

NOAA Tech. Mem. NOS ORCA 71. NOAA/NOS/ORCA, Silver Spring, MD. 85-102. 

Choi, K-S., D. H. Lewis, E. N. Powell and S. M. Ray (1993) Quantitative measurement 

of reproductive output in the American oyster, Crassostrea virginica (Gmelin), using an 

enzyme-linked immunosorbent assay (ELISA). Aquacult. Fish. Management, 

24:299-322. 

15
Choi, K-S., E. N. Powell, D. H. Lewis and S. M. Ray (1994) Instantaneous reproductive 

effort in female American oysters, Crassostrea virginica, measured by a new 

Immunoprecipitation Assay. Biol. Bull., 186:41-61. 

Cunningham, P. A. and M. R. Tripp (1975) Accumulation, tissue distribution and 

elimination of 203HgCl2 and CH3

203HgCl in the tissues of the American oyster 

Crassostrea virginica. Mar. Ecol., 31:321-34. 

Dittman, D. E., S. E. Ford and D. K. Padilla (2001) Effects of Perkinsus marinus on 

reproduction and condition of the eastern oyster, Crassostrea virginica, depend on 

timing. J. Shellfish Res., 20:1025-34. 

Ellis, M. S., R. D. Barber, R. E. Hillman, Y. Kim and E. N. Powell (1998a) 

Histopathology analysis. In: Sampling and Analytical Methods of the National Status and 

Trends Program Mussel Watch Project: 1993-1996 Update. NOAA Tech. Memo. NOS 

ORCA 130. NOAA/NOS/ORCA Silver Spring, MD. 198-215. 

Ellis, M. S., R. D. Barber, R. E. Hillman and E. N. Powell (1998b) Gonadal analysis. In: 

Sampling and Analytical Methods of the National Status and Trends Program Mussel 

Watch Project: 1993-1996 Update. NOAA Tech. Memo. NOS ORCA 130. 

NOAA/NOS/ORCA Silver Spring, MD. 216-227. 

Ford, S. E. and A. J. Figueras (1988) Effects of sublethal infection by the parasite 

Haplosporidium nelsoni (MSX) on gametogenesis, spawning, and sex ratios of oysters in 

Delaware Bay, USA. Dis. Aquat. Org., 4:121-33. 

Gurr, E. (1956) A Practical Manual of Medical and Biological Staining Techniques. 

Interscience, New York. 320 pp. 

Hillman, R. E. (1993) Gonadal index and histopathology for the East and West coasts 

used in the National Status and Trends Mussel Watch Project. In: Sampling and 

Analytical Methods of the National Status and Trends Program National Benthic 

Surveillance and Mussel Watch Projects 1984-1992. Vol. II Comprehensive descriptions 

of complementary measurements. NOAA Tech. Mem. NOS ORCA 71. 

NOAA/NOS/ORCA, Silver Spring, MD. 45-53. 

Kim, Y., E. N. Powell, T.L. Wade, B.J. Presley and J.M. Brooks (1999) Influence of 

climate change on interannual variation in contaminant body burden in Gulf of Mexico 

oysters. Mar. Environ. Res., 48:459-88. 

Kim, Y., E.N. Powell, T.L. Wade, B.J. Presley and J.M. Brooks (2001) The geographic 

distribution of population health and contaminant body burden in Gulf of Mexico oysters. 

Arch. Environ. Contam. Toxicol., 41:30-46. 

Lee, R. F. (1993) Passage of xenobiotics and their metabolites from hepatopancreas into 

ovary and oocytes of blue crabs, Callinectes sapidus: possible implications for 

vitellogenesis. Mar. Environ. Res., 35:181-7. 

16
Masson, P. (1928) Carcinoids (argentaffin cell tumors) and nerve hyperplasia of the 

appendicular mucosa. Am. J. Pathol., 4:181-211. 

Mo, C. and B. Neilson (1993) Weight and salinity effects on zinc uptake and 

accumulation for the American oyster (Crassostrea virginica Gmelin). Environ. Pollut., 

82:191-6. 

Morales-Alamo, R. and R. Mann (1989) Anatomical features in histological sections of 

Crassostrea virginica (Gmelin, 1791) as an aid in measurements of gonad area for 

reproductive assessment. J. Shellfish Res., 8:71-82. 

Newell, R. I. E. (1989) Species profiles: life histories and environmental requirements of 

coastal fishes and invertebrates (North and Mid-Atlantic) - blue mussel. U.S. Fish. Wildl. 

Serv. Biol. Rep. 82(11.102). U.S. Army Corps of Engineers, TR E1-82-4. 25 pp. 

Peters, E. C., P. P. Yevich, J. C. Harshbarger and G. E. Zaroogian (1994) Comparative 

histopathology of gonadal neoplasms in marine bivalve molluscs. Dis. Aquat. Org., 

20:59-76. 

Powell, E. N., E. A. Wilson-Ormond and K-S. Choi (1993) Gonadal analysis - 

Crassostrea virginica. In: Sampling and Analytical Methods of the National Status and 

Trends Program National Benthic Surveillance and Mussel Watch Projects 1984-1992. 

Vol. II Comprehensive descriptions of complementary measurements. NOAA Tech. 

Mem. NOS ORCA 71. NOAA/NOS/ORCA, Silver Spring, MD. 55-62. 

Preece, A. (1972) A Manual for Histologic Technicians. Little, Brown, and Company, 

Boston. 428 pp. 

Seed, R. (1975) Reproduction in Mytilus (Mollusca: Bivalvia) in European waters. 

Pubblicazioni della Stazione Zoologica di Napoli (Milan), 39(Suppl. 1):317-334. 

Seed, R. (1976) Ecology. In: Marine Mussels: Their Ecology and Physiology. B. L. 

Bayne (ed.). Cambridge University Press, New York, NY. 13-66. 

Seed, R. and T. H. Suchanek (1992) Population and community ecology of Mytilus. In: 

The Mussel Mytilus: Ecology, Physiology, Genetics and Culture. E. Gosling (Ed.). 

Developments in Aquaculture and Fisheries Science, 25:87-169. 

Sericano, J. L., T. L. Wade, E. N. Powell and J. M. Brooks (1993) Concurrent chemical 

and histological analyses: are they compatible? Chem. Ecol., 8:41-7. 

Sprung, M. (1991) Costs of reproduction: a study on metabolic requirements of the 

gonads and fecundity of the bivalve Dreissena polymorpha. Malacologia, 33:63-70. 

Supan, J. E. and C. A. Wilson (2001) Analyses of gonadal cycling by oyster broodstock, 

Crassostrea virginica (Gmelin), in Louisiana. J. Shellfish Res., 20:215-20. 

17
Wilson, E. A., E. N. Powell, M. A. Craig, T. L. Wade and J. M. Brooks (1990) The 

distribution of Perkinsus marinus in Gulf coast oysters: its relationship with temperature, 

reproduction, and pollutant body burden. Int. Rev. Gesamten Hydrobiol., 75:533-50. 

Wilson, E. A., E. N. Powell, T. L. Wade, R. J. Taylor, B. J. Presley and J. M. Brooks 

(1992) Spatial and temporal distributions of contaminant body burden and disease in Gulf 

of Mexico oyster populations: the role of local and large-scale climatic controls. Helgol. 

Meeresunters, 46:201-35. 

18
CHAPTER 2. HISTOPATHOLOGY ANALYSIS 

Y. Kim, E. N. Powell, and K. A. Ashton-Alcox 

Haskin Shellfish Research Laboratory 

Rutgers University 

Port Norris, NJ 08349 

ABSTRACT 

This chapter describes the procedures followed for histopathological analysis of oysters, 

mytilid mussels, and dreissenid mussels collected for NOAA’s National Status and 

Trends Mussel Watch Project. Analyses are conducted on paraffin-embedded tissues 

sectioned at a 5-μm thickness and stained using a pentachrome staining procedure. The 

infection intensity of parasites, the occurrence and extent of tissue pathologies, and the 

intensity of diseases are recorded using quantitative or semi-quantitative measures. 

1.0 INTRODUCTION 

The use of bivalves in the National Status and Trends (NS&T) Program is predicated 

upon their reliability as environmental integrators of contamination. The influence of 

population health on body burden and, in turn, the influence of contaminant exposure on 

population health are, as yet, poorly understood. Clearly, certain diseases (normally 

caused by viruses or single-celled prokaryotes and eukaryotes) and non-disease causing 

parasites (mostly ciliates and multi-cellular parasites) produce tissue level changes that 

might be expected to affect contaminant body burden. Gonadal quantity, for example, can 

be dramatically altered by disease (Hofmann et al., 1995; Barber, 1996; Ford and 

Figueras, 1988) and by parasites (Hopkins, 1957; Yoo and Kajihara, 1985). Certain 

contaminants are preferentially concentrated in gonadal tissue (Ellis et al., 1993; Lee, 

1993; Abbe et al., 1994). Others are concentrated in non-gonadal tissue (Mo and Neilson, 

1993; Cunningham and Tripp, 1975). Evidence, mostly by correlation, continues to 

mount for a relationship between certain tissue pathologies and contaminant exposure 

(Bowmer et al., 1991; Weis et al., 1995; Johnson et al., 1992; MacKenzie et al., 1995; 

Heinonen et al., 1999) and the influence of contaminant exposure on the bivalve immune 

system has been described (Anderson et al., 1992; Cheng, 1988; Winstead and Couch, 

1988; Ashton-Alcox et al., 2000). Besides contaminants, other environmental factors may 

facilitate disease or trigger the development of pathologies (e.g., Lee et al., 1996; 

Landsberg, 1996; Zander, 1998) with significant consequences to tissue composition and, 

in all likelihood, subsequent contaminant retention. Thus, contaminant exposure and 

health, as they feed back upon each other, may have dramatic effects on monitoring 

programs that use sentinel organisms to define spatial and temporal trends in contaminant 

loading and contaminant gradients. 

Evaluation of the health of bivalves collected as a part of the Mussel Watch Project 

necessitates determining the prevalence and intensity of diseases, parasites and 

pathologies by histological examination. Certain pathological conditions recognized by 

19
shell condition (Warburton, 1958; Lawler and Aldrich, 1987), ligament degradation 

(Dungan et al., 1989), or periostracal abnormalities (Davis and Barber, 1994) will not be 

identified using this approach and some ectoparasites are lost during collection (e.g., 

odostomians, White et al., 1985, 1989). However most common diseases, parasites, and 

pathologies can be evaluated by this method. Specific assays are available now for some 

organisms (e.g., Ko et al., 1999; Stokes and Burreson, 2001), but a histological 

examination remains the best general approach for simultaneously evaluating a wide 

range of parasites, disease and pathologies (Ford, 2002). 

Classically, histological examination involves evaluating samples for parasite prevalence 

and the occurrence of pathologies. Measures of prevalence or occurrence, however, do 

not give a true indication of the health of an organism. Wilson-Ormond et al. (2000) 

compared the usefulness of prevalence to semi-quantitative and quantitative measures of 

intensity in the Gulf of Mexico Offshore Operations Monitoring Experiment and found 

that most significant trends were observed from intensity data rather than prevalence. 

One reason for this is that prevalence depends on transmission, and transmission rate may 

be controlled by biological factors such as population density, encounter rates, and 

inherent differences in susceptibility, as well as factors acting as stressors directly on the 

individuals in the population (Kermack and McKendrick, 1991; Ackerman et al., 1984; 

Hofmann et al., 1995; Powell et al., 1996). Thus, measures of intensity or extent of tissue 

alteration may more reliably correspond to measures of exposure. Accordingly, beginning 

in 1995, a histopathological analysis designed to evaluate population health was included 

in the Mussel Watch Project. The approach taken was to evaluate the intensity of diseases 

and parasites, and the extent of tissue pathologies, rather than simply prevalence, to better 

assess the health of sampled populations. This description updates protocols described 

earlier by Ellis et al. (1998). 

A measure of overall health has not been applied, although a number of these have been 

suggested, based on tissue appearance (Quick and Mackin, 1971), histological grading 

(Bowmer et al., 1991), or summation of total parasite load (Wilson-Ormond et al., 2000). 

Although Laird (1961) argues on theoretical grounds for an underlying relationship 

between total parasite body burden and environmental quality, generally, these overall 

measures of health have not proven efficacious because the various parasites, diseases 

and pathologies originate in different ways and certain parasites, even if abundant, may 

not have a large impact on organism health. 

2.0 EQUIPMENT, REAGENTS, AND SOLUTIONS 

Preparation of samples for histopathological analysis follows the protocols established for 

gonadal analysis (see Kim et al., this volume). 

3.0 ANALYSIS 

Prepared slides are examined individually under the microscope using a 10X ocular and a 

10X objective. If any tissue needs to be examined more closely, a 25X or 40X objective 

may be used for closer examination of suspected pathologies or parasites. Major tissue 

types examined include gill, mantle, gonad and gonoducts, digestive gland tubules, 

20
stomach/digestive gland ducts, and connective tissue. Thus, a proper tissue cross-section 

is essential (Kim et al., this volume). As the histopathological analysis is done in concert 

with gonadal analysis (Kim et al., this volume), mytilid mussels are usually examined 

beginning with the gonads and mantle tissue to determine sex and stage of gonadal 

development. The gills and the visceral mass are then examined. The gonads of oysters 

and dreissenid mussels are located within/around the visceral mass, so gonads are 

examined first, followed by scanning of the visceral mass and gills. 

Parasites, diseases, or tissue pathologies are scored for intensity using either a 

quantitative or semi-quantitative scale, as described subsequently (Table 2-1). Conditions 

scored quantitatively are evaluated by keeping a running count of incidences of the 

condition as the slide is scanned to avoid re-examining each slide multiple times for each 

category. Conditions scored semi-quantitatively may require re-scanning portions of the 

tissue for each category to fully evaluate the scale of infection. 

Listed subsequently are the common parasites and pathologies encountered during 

histopathological analysis including, in each case, the method of analysis, a short 

description, and a reference to published figures of the condition. The list is not intended 

to be inclusive of all known parasites and pathologies for Mussel Watch species or all 

conditions encountered in specimens obtained during the Mussel Watch Project. 

Frequently, in routine examination, we do not attempt to differentiate at a low taxonomic 

level between related parasites because infection intensities are low for the majority of 

parasite species and so, the information gained from taxonomic analysis does not warrant 

the time spent in identification. Rather, we have lumped the various species into higher 

categories (e.g., all cestodes, all ciliates). When further differentiation is needed, we first 

differentiate by tissue of occurrence because most species have distinct tissue 

preferences. In nearly all cases, this level of differentiation has been adequate for 

estimates of prevalence and infection intensity. 

3.1. QUANTITATIVE CATEGORIES 

Most parasites are counted quantitatively (Table 2-1). These include prokaryotic 

inclusion bodies, gregarines, ciliates of various types, Pseudoklossia (a coccidian), 

cestodes, trematode metacercariae, worms ectoparasitic or commensal on the gills, 

nematodes, copepods, pinnotherid crabs, and worms in the gonoducts. We also evaluate a 

number of tissue conditions quantitatively, including the number of ceroid bodies, 

incidences of tissue inflammation, and suspected neoplasms and tumors. 

21
Table 2-1. List of quantitative and semi-quantitative categories for each bivalve taxon. In 

a number of cases, e.g., gregarines and ciliates, subcategories by tissue type and organism 

morphology are individually tallied, as described in the text. 

 Oyster Mytilid Mussel Dreissenid 

 Mussel 

Quantitative Category 

Prokaryote inclusions X X 

Gregarines X X 

Ciliates X X 

Xenomas X X 

Coccidians X 

Cestodes X 

Trematode metacercariae X X 

Turbellarians and nemerteans X X 

Nematodes X X 

Copepods X X 

Pinnotherid crabs X X 

Echinostomes X 

Unidentified organisms X X X 

Ceroid bodies X X X 

Neoplasms X 

Tissue inflammation X X X 

Tissue necrosis X X X 

Semi-quantitative Category 

Haplosporidium nelsoni (MSX) X 

Trematode sporocysts X X 

Digestive tubule atrophy X X X 

Gonadal abnormalities X X 

 (Kim et al., this volume) 

Unusual digestive tubules X X X 

________________________________________________________________________ 

22
Prokaryotic inclusions (Figures 2-1 – 2-2) [Additional photographs: Otto et al. (1979), p. 

295, Figs. 2-7; Gulka and Chang (1984), p. 320, Fig. 1; Couch (1985), p. 63, Fig. 2; 

Gauthier et al. (1990), p. 112, Fig. 7; Murchelano and MacLean (1990), p. 9, Figs. 1-5, 

1-6; Figueras et al. (1991b), p. 20, Fig. 2; Harshbarger et al. (1977), p. 667, Fig. 1; 

Robledo et al. (1994), p. 291 Fig. 3, p. 292 Fig. 4; Villalba et al. (1997), p. 130, Figs. 2-3; 

Powell et al. (1999), p. 2059 Fig. 2, p. 2060 Fig. 4], variously referred to as rickettsial 

bodies, chlamydial bodies or mycoplasms, are normally observed in the duct and tubule 

walls of the digestive gland. In Mussel Watch sites, prokaryote inclusions have been 

recorded in both mytilid mussels and oysters from the East, West, and Gulf coasts, but 

have not been observed in dreissenids from the Great Lakes area. Prokaryotic inclusions 

similar to those described by Harshbarger et al. (1977) have been observed within the 

epithelial cells of the digestive system and also occasionally occur in the lumen of the 

digestive tract in mytilids from West coast Mussel Watch sites and in oysters from East 

and Gulf coast Mussel Watch sites. In some cases, cysts containing prokaryotic 

inclusions are associated with the gill and the renal tissues in mytilids. Inclusions found 

in the digestive tract are usually roundish, whereas those in the gill and kidney are rather 

amorphous in shape. No apparent pathological effects or host responses to prokaryote 

infection have been detected, as is typical for most bivalves (Otto et al., 1979; Figueras et 

al., 1991a; Villalba et al., 1997). Each individual inclusion is counted. 

Gregarines in the genus Nematopsis (Figures 2-3 - 2-5) [Additional photographs: Cheng 

(1967), p. 148, Fig. 29; Ford (1988), p. 218, Fig. 6G; Friedman et al. (1989), p. 136, Fig. 

3; Gauthier et al. (1990), p. 110, Figs. 2, 3] are sporozoan parasites frequently found in 

oysters, and occasionally in mytilid mussels. Different Nematopsis species often show a 

tissue preference for either mantle or gill (Sprague and Orr, 1952). Kim et al. (1998) 

noted that gregarines were common in oysters from the southeastern and Gulf of Mexico 

coasts in Mussel Watch samples, and also observed gregarines in mytilid mussels from 

the West coast. Mud and stone crabs are known to be final hosts (Prytherch, 1940). 

Although gregarine infections are known to have low pathogenicity, mechanical 

interference by heavy infections has been suggested to have some harmful effects on the 

host (e.g., oyster) physiology (Sindermann, 1990). In Mussel Watch samples, gregarine 

spores typically occur in the connective tissue around the visceral mass of the body, in 

the gills, and in the mantle connective tissues of oysters and West coast mytilids. No host 

tissue reaction or discernible pathological effects have been observed, in agreement with 

Cheng (1967). Although, species identifications are not made, gregarines are routinely 

scored according to tissue occurrence; body, gill, or mantle, following Landau and 

Galtsoff (1951). Quantification is obtained by counting each individual in each tissue 

within one representative tissue section. 

23
Figure 2-1. Prokaryotic inclusions present in digestive tract epithelium of an oyster, 

Crassostrea virginica. Arrows indicate examples. x 100. 

Figure 2-2. Cyst-like encapsulations of prokaryotic microorganism in gills of a mytilid 

mussel, Mytilus edulis. x 100. 

24
Figure 2-3. Nematopsis sp. spores in connective tissue between digestive tubules of an 

oyster, C. virginica. Arrows indicate examples. x 100. 

Figure 2-4. Numerous Nematopsis sp. spores in the gills of an oyster, C. virginica. 

Arrows indicate examples. x 100. 

25
Figure 2-5. Gregarine-like spores near the tip of the mantle of a mytilid mussel, M. 

californianus. Arrows indicate examples. x 100. 

A variety of ciliate types (Figures 2-6 - 2-7) [Additional photographs: Cheng (1967), p. 

184, Fig. 88, p. 193, Fig. 102; Murchelano and MacLean (1990), p. 11 Fig. 1-11, p. 15 

Figs. 1-19, 1-20; Figueras et al. (1991a), p. 91, Fig. 2; Gauthier et al. (1990), p. 110, Figs. 

4-6; Villalba et al. (1997), p. 132, Figs. 10-11; Laruelle et al. (1999), p. 254-256, Figs. 1-

3; Moret et al. (1999), p. 36, Fig. 1] have been observed in bivalves from Mussel Watch 

sites (Kim et al., 1998), normally at low prevalence and more frequently in mytilid 

mussels than in oysters. Ciliate infections are observed in mytilids from the East and 

West coasts and in oysters from the East and Gulf coasts. Gill ciliates are one of the most 

commonly observed parasites. Ciliates occur in between gill filaments or are attached to 

gill surfaces of mytilids and oysters. Ciliates are also found in the gut lumen or attached 

to the digestive tract epithelia. Otto et al. (1979), Figueras et al. (1991a) and Villalba et 

al. (1997) reported no notable pathology in bivalves parasitized by ciliates and ciliate 

infections do not appear to elicit any obvious pathological conditions or host responses in 

Mussel Watch samples. Ciliates are quantitated by tissue type (e.g., gut, digestive gland, 

gill). Xenomas (Figure 2-8), cells distended with maturing ciliates, are tabulated 

separately. If a large xenoma has burst, the individual small ciliates are counted. 

26
Figure 2-6. Ciliates between the gill filaments of a mytilid mussel, M. edulis. Arrows 

indicate examples. x 100. 

Coccidians of the genus Pseudoklossia (Apicomplexa) [Photographs: Morado et al. 

(1984), p. 212 Figs. 7-10, p. 213 Figs. 11-16; Friedman et al. (1995), p. 35, Figs. 3-11; 

Villalba et al. (1997), p. 130, Figs. 5-6] are another protozoan parasite and are 

occasionally observed in the kidney of mytilid mussels at Mussel Watch sites. Each 

parasite is counted. 

A variety of encysted larval cestodes (Figures 2-9 – 2-10) [Additional photographs: 

Cheng (1966a), p. 248 Fig. 6, p. 252 Figs. 1-6, p. 254 Figs. 1-2; Murchelano and 

MacLean (1990), p. 17, Fig. 1-21; Sindermann (1970), p. 128, Fig. 42] (for examples, see 

Cake, 1977; Cake and Menzel, 1980) have been observed. Encysted cestodes have been 

observed in either connective tissue around the digestive gland and gut or in the gills of 

oysters in Mussel Watch samples. None have been observed in mytilid mussels or 

dreissenids. From histological examination of their tissue location in oysters, cestodes 

presumably penetrate the gill or digestive epithelium of the host bivalve. Cestode 

infection does not seem to significantly damage the oyster. Cellular reaction to cestode 

cysts, characterized by encapsulation of larval cestodes by layers of connective tissue 

fibers (Cheng, 1966a; Sindermann, 1970), is routinely observed. Encapsulated larval 

cestodes normally appear to be disintegrating and to be in the process of resorption. 

Cestodes are quantified by tissue location (e.g., body, gill, mantle). Each occurrence 

observed is counted separately. 

27
Figure 2-7. Ciliates in the lumen and attached to the epithelium of the intestine of an 

oyster, C. virginica. Arrows indicate examples. x 100. 

Figure 2-8. Ciliated xenomas on the gill surface of an oyster, C. virginica. Arrows 

indicate examples. x 100. 

28
Metacercariae of trematodes (nearly all Proctoeces sp.) [Photographs: Little et al. (1969), 

p. 455, Fig. 1; Wolf et al. (1987), p. 380, Fig. 1; Tripp and Turner (1978), p. 77 Fig. 4, p. 

79 Figs. 5-8, p. 81 Figs. 9-12; Winstead and Couch (1981), p. 297 Fig. 1, p. 299 Figs. 2-

3] have been observed, normally at low prevalence, and occur in the mantle, foot, 

gonad/gonoduct and pericardial cavity of mytilid mussels and in the gonoduct of oysters 

at Mussel Watch sites. Proctoeces and occasional nematodes also found in the gonoduct 

should be distinguished from the echinostome metacercariae [Photograph: Ellis et al. 

(1998), p. 201, Fig. 1] observed in the gonoducts of oysters from the Gulf of Mexico 

(Winstead et al., 1998). Encysted metacercariae (Figure 2-11) (presumably 

gymnophallids) [Additional photograph: Bower et al. (1994), p. 74, Fig. 62] are 

frequently observed in all tissues of mytilids: mantle, visceral connective tissue, foot, 

byssal gland and gill. In most cases, no conspicuous host response is observed. 

Hemocytes, however, occasionally infiltrate and surround the worms, especially those 

that are dead or dying. Each trematode occurrence is counted separately. 

Gill nemerteans and turbellarians (Brun et al., 1999) [Photographs: Villalba et al. (1997), 

p. 132, Figs. 12-13; Cáceres-Martínez et al. (1998), p. 218, Figs. 3-4] are occasionally 

seen between gill filaments. Whether these are commensal or ectoparasitic is unknown. 

Each cross-section observed is counted although one individual may be responsible for a 

number of tissue cross-sections. However, this method of quantification has proven 

effective even at high infection intensities (Wilson-Ormond et al., 2000). 

Figure 2-9. Encapsulated larval cestodes in the vesicular connective tissue surrounding 

the stomach of an oyster, C. virginica. x 63. 

29
Figure 2-10. A larval cestode in the gill connective tissue of an oyster, C. virginica. x 

100. 

Nematodes (Figure 2-12) [Additional photographs: Cheng (1967), p. 264, Fig. 172; Lowe 

and Moore (1979), p. 140, Fig. 8; Gauthier et al. (1990), p. 112, Fig. 9; Murchelano and 

MacLean (1990), p. 19, Fig. 1-25; Sparks (1985), p. 375-376, Figs. 10-16] are 

occasionally observed parasitizing oysters from the East and Gulf coasts, and also 

dreissenids in Mussel Watch samples (Kim et al., 1998). Nematodes reported in molluscs 

are usually larval stages (Cheng 1978; Lichtenfels et al., 1980). Adults are found in 

predators of molluscs (Cheng, 1978), such as elasmobranchs (Millemann, 1963) and sea 

turtles (Berry and Cannon, 1981). Cheng (1966b) suggested that larval nematodes invade 

oysters via the digestive tract and migrate through tissues by way of blood vessels. In 

Mussel Watch samples, larval nematodes in oysters have been found localized in 

vesicular connective tissues around the region of the digestive gland, as described by 

Burton (1963) and Couch (1985), destroying adjacent host tissues. In some cases, a host 

cellular response, infiltration of hemocytes, is observed in association with the worm, as 

was reported by Couch (1985). Each individual cross-section is counted separately, 

although, like the nemerteans, a single individual may be responsible for a number of 

cross-sections. 

Parasitic copepods [Photographs: Lowe and Moore (1979), p. 140, Fig. 7; Murchelano 

and MacLean (1990), p. 19, Fig. 1-26; Figueras et al. (1991b), p. 27, Fig. 6; Robledo et 

al. (1994), p. 295 Fig. 8; Villalba et al. (1997), p. 132, Fig. 15] are occasionally found in 

the gut lumen (Gee and Davey, 1986). Each occurrence is counted. 

30
Figure 2-11. Cross-sections of metacercariae encysted in the visceral connective tissue of 

a mytilid mussel, M. edulis. x 63. 

Figure 2-12. Sections of unidentified nematode larvae in the digestive gland connective 

tissue of an oyster, C. virginica. Arrows indicate examples. x 100. 

31
Pinnotherid crabs [Photograph: Stauber (1945), p. 277, Fig. 23; Sandoz and Hopkins 

(1947), p. 257, Plate III] are occasionally found in the mantle cavity of oysters and 

mytilid mussels at Mussel Watch sites. Gill damage in infected hosts (Stauber, 1945; 

Christensen and McDermott, 1958; Haven, 1959) is frequently observed. Pinnotherid 

crabs also deprive the host of food (Stauber, 1945; Bierbaum and Shumway, 1988). Each 

occurrence is counted. 

Ceroid bodies or brown cells [Photograph: Cheng and Burton (1965), p. 6, Fig. 5; Farley 

(1968), p. 590 Fig. 16, p. 592 Fig. 23; Murchelano and MacLean (1990), p. 11, Figs. 1-9, 

1-12] are distinct brown-yellow aggregates that may occur in large clumps, and appear to 

be involved in metabolite accumulation and detoxification (Zaroogian and Yevich, 1993). 

Typically, they occur in greatest abundance in oysters, and in lesser numbers in mytilid 

mussels and dreissenid mussels. Quantification is obtained by counting each ceroid body. 

Occasionally, a ceroid body appears fractured or split; in this case only one fragment is 

counted. 

Tumors and neoplasms (Figure 2-13) [Additional photographs: Murchelano and 

MacLean (1990), p. 19, Figs. 1-27, 1-28; Peters (1988), p. 81, Figs. C, E, F; Sparks 

(1985), p. 107 Fig. 27, p. 113 Fig. 42; Couch (1985), p. 69 Fig. 10, p. 70 Fig. 11; 

Figueras et al. (1991b), p. 30, Fig. 10; Villalba et al. (1997), p. 132, Fig. 16] are 

occasionally observed. Disseminated sarcomas, probably of hematopoietic origin, are 

particularly common in mytilid mussels in the Puget Sound region (e.g., Elston et al., 

1990). The occurrence of neoplasms and tumors in oysters is extremely rare. Examples 

are described by Farley (1969, 1976), Harshbarger et al. (1979) and Ford and Tripp 

(1996). Neoplasms are occasionally observed in mytilids in Mussel Watch samples. 

Neoplastic cells with characteristic high nucleus-to-cytoplasm ratios (Ford et al., 1997) 

fill the vesicular connective tissues of the affected mytilids. All observed disseminated 

sarcomas have been seen in mytilids. For each specimen examined, neoplasms are 

recorded as either present or absent. 

Cases of tissue inflammation (Figures 2-14 – 2-15) [Additional photographs: Farley 

(1968), p. 590, Fig. 17; Couch (1985), p. 65, Fig. 5; Figueras et al. (1991b), p. 28 Fig. 7, 

p. 29 Fig. 8; Murchelano and MacLean (1990), p. 11, Figs. 1-9, 1-10; Lowe and Moore 

(1979), p. 138, Figs. 1-4; Villalba et al. (1997), p. 132, Fig. 17; Sindermann (1970), p. 

110, Fig. 34] characterized by intense infiltration of hemocytes may be focal or diffuse. 

The type of affected tissue and type of irritation responsible influences the nature of the 

cellular response (Ford and Tripp, 1996). Diffuse inflammation is differentiated from 

focal inflammation when the affected area does not appear to have a clear center or focal 

point of highest hemocyte concentration and hemocytes are abundant and distributed 

broadly over a large section of tissue. In Mussel Watch samples, most tissue 

inflammation, characterized by hemocytic infiltration, and most tissue necrosis, 

characterized by death or decay of cells and tissues, is observed in the visceral connective 

tissue and is sometimes associated with the presence of parasites. Granulocytomas 

(Figure 2-15), an inflammatory cellular condition characterized by clusters of hemocytes 

or the disintegration and sloughing of tissue, occur mainly in the digestive gland of 

mytilids, as was reported by Villalba et al. (1997). These tissue pathologies, focal 

32
inflammation, diffuse inflammation and tissue necrosis, are tallied separately. Each 

affected area is counted. 

3.2. SEMI-QUANTITATIVE CATEGORIES 

Some conditions are assigned to a semi-quantitative scale related to the intensity or the 

extent of the affected area (Table 2-1). With one exception, so-called digestive gland 

atrophy, these are pathologies affecting large tissue areas, diseases characterized by 

systemic effects, or parasites for which individual counts are not feasible. Semiquantitative categories include Haplosporidium nelsoni (MSX), trematode sporocysts, 

unusual digestive tubules, gonadal abnormalities (discussed in Kim et al., this volume), 

and digestive gland atrophy. Perkinsus marinus, an oyster parasite that is also assayed 

semi-quantitatively, is assayed by the more precise thioglycollate method, rather than by 

histology (Ashton-Alcox et al., this volume). 

Figure 2-13. Neoplastic cells infiltrating the visceral connective tissue of a mytilid 

mussel, M. edulis. x 100. 

Haplosporidium nelsoni (Figure 2-16) [Additional photographs: Farley (1968), p. 590 

Fig. 13, p. 592 Fig. 21; Ford (1988), p. 214, Fig. 4; Ford and Tripp (1996), p. 617, Fig. 

20], the haplosporidan protozoan responsible for MSX (multinucleated sphere X) disease 

in eastern oysters, was first reported in Delaware Bay oysters in the late 1950s (Haskin et 

al., 1965). Haplosporidium nelsoni was likely introduced from Japan (Burreson et al., 

2000). It now ranges from Maine to Florida along the entire East coast (Kern, 1988; Ford 

and Tripp, 1996). Kim et al. (1998) observed Haplosporidium nelsoni in oysters from 

Delaware Bay to Georgia at Mussel Watch sites. Multinucleated plasmodia are observed 

in epithelial cells and the connective tissues of the gills and digestive tract. 

33
Figure 2-14. Hemocytic infiltration near the gill base of an oyster, C. virginica. Arrows 

indicate examples. x 100. 

Figure 2-15. Granulocytomas in the digestive gland of a mytilid mussel, M. edulis. 

Arrows indicate examples. x 40. 

Haplosporidium nelsoni infections start in the gill epithelium and are limited to this area 

at light infection levels. As the disease worsens, it becomes systemic and is eventually 

found throughout the visceral mass. MSX disease, especially heavy infections, is 

34
associated with host hemocyte infiltration into the site of infection and tissue necrosis as 

observed by Farley (1968) and Ford (1985). 

Because of the small size of Haplosporidium nelsoni, oyster tissues may need to be 

examined at a higher power than 10X. MSX infection normally starts in the gill 

epithelium so the gill tissue must be carefully examined to score early infections 

accurately. In cases where the disease has become systemic, examination of the visceral 

mass is necessary to score infection intensity. Grading MSX infection is a two-step 

process. First, the intensity of the infection is graded according to a semi-quantitative 

scale (Table 2-2) that records the location, parasite numbers, epithelial or systemic, and 

extent of infection in the gill tissue or the body tissue separately. In the second step, the 

two separate ratings are composited into a 0-4 scale (Table 2-3). To get the composite 

rating, the semi-quantitative rankings for the gill and the body are applied to the matrix in 

Table 2-2. Semi-quantitative scale for Haplosporidium nelsoni infection modified from 

Ford (1985, 1986), and Ford and Figueras (1988). 

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 systemic 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 immersion field 

6 Heavy systemic infection, averaging > 20 parasites per 1000X oil 

immersion field 

35
Table 2-3. Composite rating matrix for Haplosporidium nelsoni infection. 

Composite rating scheme 

Body 

0 1 2 3 4 5 6 

0 0 1 1 2 

1 1 1 1 2 

2 1 1 1 2 2 

Gill 3 2 2 2 3 3 

4 2 2 3 3 3 4 

5 2 3 3 3 4 4 4 

6 3 4 4 4 

Figure 2-16. Numerous multinucleated plasmodia of Haplosporidium nelsoni in the gills 

of an oyster, C. virginica. Arrows point to example parasites. x 100. 

36
Trematode sporocysts of the families Fellodistomidae and Bucephalidae (Figures 2-17 – 

2-18) [Additional photographs: Ellis et al. (1998), p. 207-208, Figs. 7-9; Cheng and 

Burton (1965), p. 6 Figs. 1-4, p. 8 Figs. 10-12, p. 10 Figs. 14-17; Tripp and Turner 

(1978), p. 77, Figs. 1-3; Gauthier et al. (1990), p. 112, Fig. 8; Murchelano and MacLean 

(1990), p. 17, Fig. 1-23; Sindermann (1970), p. 125, Fig. 41; Figueras et al. (1991a), p. 

92, Fig. 3; Davids and Kraak (1993), p. 751, Fig. 1; Robledo et al. (1994), p. 294, Fig. 7; 

Villalba et al. (1997), p. 132, Fig. 14; Powell et al. (1999), p. 2061 Fig. 5] occur 

principally in the gonadal tissue of oysters (Hopkins, 1957) and mytilid mussels. Kim et 

al. (1998) reported trematode sporocyst infections in East and West coast mytilid mussels 

and in Gulf oysters from Mussel Watch sites. Carnivorous fish are the final host of 

bucephalid trematodes (Hopkins, 1954). Fellodistomid trematodes of the genus 

Proctoeces can complete their entire life cycle in a single invertebrate host (e.g., mytilid 

mussels). Thus, they have a unique life cycle involving molluscs as regular final hosts 

and bottom fishes as alternative final or as post-cycle hosts (Stunkard and Uzmann, 

1959). In Mussel Watch samples, trematodes that have invasive and ramifying sporocysts 

occur principally in the visceral connective tissues of the digestive gland and the gonadal 

tissue, destroying gametic tissue and often causing host sterilization. Sindermann (1990) 

noted that sterilization and tissue destruction are the principal result of the sporocyst 

invasion. Sterilization is normally observed in infected individuals in Mussel Watch 

samples (Hillman et al., 1988). In extensive and advanced stages of infection, sporocysts 

infiltrate the gill, mantle and other tissues (see also Cheng and Burton, 1965). Little or no 

apparent aggregation of host hemocytes around healthy sporocysts and no other evident 

host reactions have been observed in Mussel Watch samples, which concurs with Cheng 

and Burton (1965). However, infiltration of hemocytes is occasionally observed around 

dead or degenerating parasites in Mussel Watch samples, as was reported by Teia dos 

Santos and Coimbra (1995). The large branching sporocysts are difficult to quantify. 

Hence, infection intensity is scored on a semi-quantitative scale (Table 2-4). Pictorial 

examples of the rating scale include Figure 2-17 (scored 1) and Figure 2-18 (scored 2). 

37
Table 2-4. Semi-quantitative scale for trematode sporocyst infection. 

Score Description 

0 Uninfected 

1 Present in the gonads only (some gametic tissue still present) 

2 Completely filling the gonads (no gametic tissue present); may be 

present in digestive 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 gonad; substantially filling the digestive 

gland or gill; individuals appear to be a sac of sporocyst 

Figure 2-17. This trematode sporocyst infection in Mytilus edulis scored a 1 according to 

Table 2-4. Some gametic tissue is still present. 

38
Figure 2-18. This trematode sporocyst infection in Mytilus edulis scored a 2 according to 

Table 2-4. No gametic tissue is present. 

Digestive gland atrophy (Figures 2-19 – 2-21 reproduced from Figures 4-6 in Ellis et al., 

1998) [Additional photographs: Ellis et al. (1998), p. 205-206, Figs. 4-6; Couch (1985), 

p. 66, Fig. 6; Gauthier et al. (1990), p. 112, Fig. 10; Winstead (1995), p. 107, Figs. 3-4], a 

condition characterized by the thinning of the digestive tubule walls, has been described 

in a number of bivalve species (Bielefeld, 1991; Marigómez et al., 1990; Axiak et al., 

1988). Causes of the condition have been ascribed to a variety of stressors including 

exposure to contaminants and variations in food supply. Winstead (1995) found that poor 

nutrition was a key element in producing the condition in oysters and that the digestive 

gland recovered to its normal state relatively rapidly once food supply improved. It is, 

therefore, not necessarily a pathology. The digestive gland is scanned for tubules showing 

evidence of epithelial thinning. The average degree of thinning is assigned a numerical 

rating (Table 2-5). The semi-quantitative assessment permits the reading of many 

samples in a short time. For increased accuracy, the ratio of tubule diameter to wall 

diameter (e.g., Winstead, 1995) or a direct measure of wall thickness (Marigómez et al., 

1990) can be used. 

39
Table 2-5. 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 

thickness, 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 

atrophied, some walls extremely thin (fully atrophied) 

4 Wall extremely thin (100% atrophied), nearly all tubules affected 

Figure 2-19. Crassostrea virginica normal digestive tubule, scored a 0 according to Table 

2-5. 

40
Figure 2-20. Digestive gland atrophy in Crassostrea virginica scored a 2 according to 

Table 2-5. 

Figure 2-21. Digestive gland atrophy in Crassostrea virginica scored a 4 according to 

Table 2-5. 

41
In Mussel Watch samples, degenerated and/or necrotic digestive glands (Figure 2-22) 

[Additional photograph: Couch (1985), p. 67, Fig. 7] were frequently observed, 

particularly in mytilids. This condition is characterized by digestive tubules in unusually 

poor condition with loss of their normal integrity and structure, and sometimes with 

vacuolated epithelium. Individual digestive tubules are sometimes not discernible from 

each other. For each specimen examined, unusual digestive tubules are recorded as either 

present or absent. 

3.3. SUMMARY STATISTICS 

Three descriptions of parasite distribution are used: prevalence, infection intensity and 

weighted prevalence (Ford, 1988). Prevalence describes the proportion of individuals in 

the population that are infected by a specific parasite or pathology and is calculated as: 

 number of hosts with parasite or pathology 

prevalence = ______________________________________________________ 

number of hosts analyzed 

Infection intensity is calculated as the average number of occurrences of the parasite or 

pathology in infected hosts. This is a measure of the intensity of infection in infected 

individuals. 

 total number of occurrences of parasite or pathology 

infection intensity = __________________________________________________________________ 

number of hosts with parasite or pathology 

Weighted prevalence or mean abundance (Bush et al., 1997; Rózsa et al., 2000) is the 

multiple of prevalence and infection intensity, and is a measure of the relative severity of 

infection within the population. Weighted prevalence is calculated as: 

 total number of occurrences of parasite or pathology 

weighted prevalence = __________________________________________________________________ 

number of hosts analyzed 

42
Figure 2-22. Unusual digestive tubules of a mytilid mussel, M. edulis. Note the absence 

of normal tubule structure as shown in Figures 2-20 and 2-21. x 100. 

4.0 CONCLUSIONS 

The described techniques provide the quantitative and semi-quantitative methods used to 

determine the prevalence and infection intensity of parasites, pathologies, and diseases 

affecting oysters, mytilid mussels, and dreissenid mussels in the Mussel Watch Project. 

The described histopathological method is an approach that targets a wide range of 

parasites and pathologies. Specific conditions are often better assessed by other methods 

[e.g., Perkinsus marinus infection in oysters (Ashton-Alcox et al., this volume)]. The 

described method emphasizes the quantification of infection intensity. Prevalence rarely 

provides an adequate description of the population dynamics of disease and, in practice, 

often yields ambiguous results. Infection intensity as quantified by direct counts or the 

use of semi-quantitative scales consistently provides a more robust data set for statistical 

analysis comparing the spatial and temporal distribution of parasites, pathologies, and 

diseases to contaminant body burden. 

5.0 REFERENCES 

Abbe, G. R., J. G. Sanders and G. F. Riedel (1994) Silver uptake by the oyster 

(Crassostrea virginica): effect of organism size and storage sites. Estuarine Coastal Shelf 

Sci., 39:249-60. 

Ackerman, E., L. R. Elveback and J. P. Fox (1984) Simulation of Infectious Disease 

Epidemics. Charles C. Thomas, Springfield, IL. 202 pp. 

43
Anderson, R. S., L. M. Oliver and D. Jacobs (1992) Immunotoxicity of cadmium for the 

eastern oyster (Crassostrea virginica [Gmelin, 1791]): effects of hemocyte 

chemiluminescence. J. Shellfish Res., 11: 31-5. 

Ashton-Alcox, K. A., B. Allam and S. E. Ford (2000) Application of flow cytometry to 

bivalve pathology. In: Recent Advances in Marine Biotechnology. Vol. 5 

Immunobiology and pathology. M. Fingerman and R. Nagabhushanam (eds.). Science 

Publishers, Inc., New Hampshire. 85-124. 

Axiak, V., J. J. George and M. N. Moore (1988) Petroleum hydrocarbons in the marine 

bivalve Venus verrucosa: accumulation and cellular responses. Mar. Biol. (Berl.), 

97:225-30. 

Barber, B. J. (1996) Gametogenesis of eastern oysters, Crassostrea virginica (Gmelin, 

1791), and Pacific oysters, Crassostrea gigas (Thunberg, 1793) in disease-endemic lower 

Chesapeake Bay. J. Shellfish Res., 15:285-90. 

Berry, G. N. and L. R. G. Cannon (1981) The life history of Sulcascaris sulcata

(Nematoda: Ascaridoidea), a parasite of marine molluscs and turtles. Intl. J. Parasitol., 

11:43-54. 

Bielefeld, U. (1991) Histological observation of gonads and digestive gland in starving 

Dreissena polymorpha (Bivalvia). Malacologia, 33:31-42. 

Bierbaum, R. and S. E. Shumway (1988) Filtration and oxygen consumption in mussels, 

Mytilus edulis, with and without pea crabs, Pinnotheres maculatus. Estuaries, 11:264-71. 

Bower, S. M., S. E. McGladdery and I. M. Price (1994) Synopsis of infectious diseases 

and parasites of commercially exploited shellfish. Ann. Rev. Fish Dis., 4:1-199. 

Bowmer, C. T., M. van der Meer and M. C. T. Scholten (1991) A histopathological 

analysis of wild and transplanted Dreissena polymorpha from the Dutch sector of the 

River Maas. Comp. Biochem. Physiol. C, 100:225-9. 

Brun, N. T., A. D. Boghen and J. Allard (1999) Distribution of the turbellarian Urastoma 

cyprinae on the gills of the eastern oyster Crassostrea virginica. J. Shellfish Res., 

18:175-9. 

Burreson, E. M., N. A. Stokes and C. S. Friedman (2000) Increased virulence in an 

introduced pathogen: Haplosporidium nelsoni (MSX) in the eastern oyster Crassostrea 

virginica. J. Aquat. Animal Health, 12:1-8. 

Burton, R. W. (1963) Distribution of oyster microparasites in Chesapeake Bay, 

Maryland, 1959-1960. Proc. Natl. Shellfish. Assoc., 52:65-74. 

Bush, A. O., K. D. Lafferty, J. M. Lotz and A. W. Shostak (1997) Parasitology meets 

ecology on its own terms: Margolis et al. revisited. J. Parasitol., 83:575-83. 

44
Cáceres-Martínez, J., R. Vásquez-Yeomans and R. Sluys (1998) The turbellarian 

Urastoma cyprinae from edible mussels Mytilus galloprovincialis and Mytilus 

californianus in Baja California, NW Mexico. J. Invertebr. Pathol., 72:214–9. 

Cake Jr., E. W. (1977) Larval cestode parasites of edible mollusks of the northeastern 

Gulf of Mexico. Gulf. Res. Rpt., 6:1-8. 

Cake Jr., E. W. and R. W. Menzel (1980) Infections of Tylocephalum metacestodes in 

commercial oysters and three predaceous gastropods of the eastern Gulf of Mexico. Proc. 

Natl. Shellfish. Assoc., 70:94-104. 

Cheng, T. C. (1966a) The coracidium of the cestode Tylocephalum and the migration and 

fate of this parasite in the American oyster, Crassostrea virginica. Trans. Amer. Microsc. 

Soc., 85:246-55. 

Cheng, T. C. (1966b) Perivascular leucocytosis and other types of cellular reactions in the 

oyster Crassostrea virginica experimentally infected with the nematode Angiostrongylus 

cantonensis. J. Invertebr. Pathol., 8:52-8. 

Cheng, T. C. (1967) Marine molluscs as hosts for symbioses with a review of known 

parasites of commercially important species. Adv. Mar. Biol., Vol. 5, Academic Press, 

London. 424 pp. 

Cheng, T. C. (1978) Larval nematodes parasitic in shellfish. Mar. Fish. Rev., 40:39-42. 

Cheng, T. C. (1988) In vivo effects of heavy metals on cellular defense mechanisms of 

Crassostrea virginica: total and differential cell counts. J. Invertebr. Pathol., 51:207-14. 

Cheng, T. C. and R. W. Burton (1965) Relationships between Bucephalus sp. and 

Crassostrea virginica: histopathology and sites of infection. Chesapeake Sci., 6:3-16. 

Christensen, A. M. and J. J. McDermott (1958) Life-history and biology of the oyster 

crab, Pinnotheres ostreum Say. Biol. Bull., (Woods Hole) 114:146-79. 

Couch, J. A. (1985) Prospective study of infectious and noninfectious diseases in oysters 

and fishes in three Gulf of Mexico estuaries. Dis. Aquat. Org., 1:59-82. 

Cunningham, P. A. and M. R. Tripp (1975) Accumulation, tissue distribution and 

elimination of 203HgCl2 and CH3

203HgCl in the tissues of the American oyster 

Crassostrea virginica. Mar. Ecol., 31:321-34. 

Davids, C. and M. H. S. Kraak (1993) Trematode parasites of the zebra mussel 

(Dreissena polymorpha). In: Zebra Mussels: Biology, Impacts, and Control. T. F. Nalepa 

and D. W. Schloesser (eds.). Lewis Publishers, Boca Raton, FL. 749-60. 

45
Davis, C. V. and B. J. Barber (1994) Size-dependent mortality in hatchery reared 

populations of oysters, Crassostrea virginica, Gmelin 1791, affected by juvenile oyster 

disease. J. Shellfish Res., 13:137-42. 

Dungan, C. F., R. A. Elston and M. H. Schiewe (1989) Evidence for colonization and 

destruction of hinge ligaments in cultured juvenile Pacific oysters (Crassostrea gigas) by 

cytophaga-like bacteria. Appl. Environ. Microbiol., 55:1128-35. 

Ellis, M. S., R. D. Barber, R. E. Hillman, Y. Kim and E. N. Powell (1998) 

Histopathology analysis. In: Sampling and Analytical Methods of the National Status and 

Trends Program Mussel Watch Projects: 1993-1996 Update. NOAA Tech. Memo. NOS 

ORCA 130. NOAA/NOS/ORCA Silver Spring, MD. 198-215. 

Ellis, M. S., K-S. Choi, T. L. Wade, E. N. Powell, T. J. Jackson and D. H. Lewis (1993) 

Sources of local variation in polynuclear aromatic hydrocarbon and pesticide body 

burden in oysters (Crassostrea virginica) from Galveston Bay, Texas. Comp. Biochem. 

Physiol. C, 106:689-98. 

Elston, R. A., A. S. Drum and S. K. Allen, Jr. (1990) Progressive development of 

circulating polyploid cells in Mytilus with hemic neoplasia. Dis. Aquat. Org., 8:51-9. 

Farley, C. A. (1968) Minchinia nelsoni (Haplosporida) disease syndrome in the American 

oyster Crassostrea virginica. J. Protozool., 15:585-99. 

Farley, C. A. (1969) Probable neoplastic disease of the hematopoietic system in oysters, 

Crassostrea virginica and Crassostrea gigas. Natl. Cancer Inst. Monogr., 31:541-55. 

Farley, C. A. (1976) Proliferative disorders in bivalve mollusks. Mar. Fish. Rev., 38:30-3. 

Figueras, A. J., C. F. Jardon and J. R. Caldas (1991a) Diseases and parasites of mussels 

(Mytilus edulis, Linnaeus, 1758) from two sites on the east coast of the United States. J. 

Shellfish Res., 10:89-94. 

Figueras, A. J., C. F. Jardon and J. R. Caldas (1991b) Diseases and parasites of rafted 

mussels (Mytilus galloprovincialis Lmk): preliminary results. Aquaculture, 99:17-33. 

Ford, S. E. (1985) Chronic infections of Haplosporidium nelsoni (MSX) in the oyster 

Crassostrea virginica. J. Invertebr. Pathol., 45:94-107. 

Ford, S. E. (1986) Comparison of hemolymph proteins from resistant and susceptible 

oysters, Crassostrea virginica, exposed to the parasite Haplosporidium nelsoni (MSX). J. 

Invertebr. Pathol., 47:283-94. 

Ford, S. E. (1988) Host-parasite interactions in eastern oysters selected for resistance to 

Haplosporidium nelsoni (MSX) disease: survival mechanisms against a natural pathogen. 

In: Disease Processes in Marine Bivalve Molluscs. W. S. Fisher (ed.). Am. Fish. Soc. 

Spec. Publ., 18:206-24. 

46
Ford, S. (2002) Testing for molluscan pathogens: an overview. In: Eastern United States 

Interstate Shellfish Seed Transport Workshop. W. D. Anderson, C. Battey, M. R. DeVoe, 

J. N. Kraeuter, A. Segars and J. M. Whetstone (eds.). 6-9. 

Ford, S. E., R. D. Barber and E. Marks (1997) Disseminated neoplasia in juvenile eastern 

oysters Crassostrea virginica, and its relationship to the reproductive cycle. Dis. Aquat. 

Org., 28:73-7. 

Ford, S. E. and A. J. Figueras (1988) Effects of sublethal infection by the parasite 

Haplosporidium nelsoni (MSX) on gametogenesis, spawning, and sex ratios of oysters in 

Delaware Bay, USA. Dis. Aquat. Org., 4:121-33. 

Ford, S. E. and M. R. Tripp (1996) Diseases and defense mechanisms. In: The Eastern 

Oyster Crassostrea virginica. V. S. Kennedy, R. I. E. Newell and A. F. Eble (eds.). 

Maryland Sea Grant College, University of Maryland, College Park, MD. 581-660. 

Friedman, C. S., G. R. Gardner, R. P. Hedrick, M. Stephenson, R. J. Cawthorn and S. J. 

Upton (1995) Pseudoklossia haliotis sp.n. (Apicomplexa) from the kidney of California 

abalone, Haliotis spp. (Mollusca). J. Invertebr. Pathol., 66:33-8. 

Friedman, C. S., T. McDowell, J. M. Groff, J. T. Hollibaugh, D. Manzer and R. P. 

Hedrick (1989) Presence of Bonamia ostreae among populations of the European flat 

oyster, Ostrea edulis Linnaeus, 1750, in California, USA. J. Shellfish Res., 8:133-7. 

Gauthier, J. D., T. M. Soniat and J. S. Rogers (1990) A parasitological survey of oysters 

along salinity gradients in coastal Louisiana. J. World Aquaculture Soc., 21:105-15. 

Gee, J. M. and J. T. Davey (1986) Experimental studies on the infestation of Mytilus 

edulis (L.) by Mytilicola intestinalis Steuer (Copepoda, Cyclopoida). J. Cons. Int. Explor. 

Mer., 42:265-71. 

Gulka, G. and P. W. Chang (1984) Host response to rickettsial infection in blue mussel, 

Mytilus edulis L. J. Fish Dis., 8:319-23. 

Harshbarger, J. C., S. C. Chang and S. V. Otto (1977) Chlamydiae (with phages), 

mycoplasmas, and rickettsiae in Chesapeake Bay bivalves. Science, 196:666-8. 

Harshbarger, J. C., S. V. Otto and S. C. Chang (1979) Proliferative disorders in 

Crassostrea virginica and Mya arenaria from the Chesapeake Bay and intranuclear 

virus-like inclusions in Mya arenaria with germinomas from a Maine oil spill site. 

Haliotis, 8:243-8. 

Haskin, H. H., W. J. Canzonier and J. L. Myhre (1965) The history of MSX on Delaware 

Bay oyster grounds, 1957-65. Amer. Malacol. Union Bull., 32:20-1. 

Haven, D. (1959) Effects of pea crabs Pinnotheres ostreum on oysters Crassostrea 

virginica. Proc. Natl. Shellfish. Assoc., 49:77-86. 

47
Heinonen, J., J. V. K. Kukkonen and I. J. Holopainen (1999) The effects of parasites and 

temperature on the accumulation of xenobiotics in a freshwater clam. Ecol. Appl., 9:475-

81. 

Hillman, R. E., P. D. Boehm and S. Y. Freitas (1988) A pathology potpourri from the 

NOAA Mussel Watch Project. J. Shellfish Res., 7:216-7. 

Hofmann, E. E., E. N. Powell, J. M. Klinck and G. Saunders (1995) Modeling diseased 

oyster populations I. Modelling Perkinsus marinus infections in oysters. J. Shellfish Res., 

14:121-51. 

Hopkins, S. H. (1954) The American species of trematode confused with Bucephalus

(Bucephlopsis) haimeanus. Parasitology, 44:353-70. 

Hopkins, S. H. (1957) Our present knowledge of the oyster parasite “Bucephalus”. Proc. 

Natl. Shellfish. Assoc., 47:58-61. 

Johnson, L. L., C. M. Stehr, O. P. Olson, M. S. Myers, S. M. Pierce, B. B. McCain and 

U. Varanasi (1992) National Benthic Surveillance Project: Northeast Coast. Fish 

histopathology and relationships between lesions and chemical contaminants (1987-89). 

NOAA Tech. Memo. NMFS-NWFSC-4. NOAA/NMFS, Seattle, WA. 95 pp. 

Kermack, W. O. and A. G. McKendrick (1991) Contributions to the mathematical theory 

of epidemics. - I. Bull. Math. Biol., 53:33-55. 

Kern, F. G. (1988) Recent changes in the range of “MSX” Haplosporidium nelsoni. J.

Shellfish Res., 7:543-4. 

Kim, Y., E.N. Powell, T.L. Wade, B.J. Presley and J. Sericano (1998) Parasites of 

sentinel bivalves in the NOAA Status and Trends Program: distribution and relationship 

to contaminant body burden. Mar. Poll. Bull., 37:45-55. 

Ko, Y-T., M. M-Y. Chan, S. E. Ford and D. Fong (1999) A PCR-ELISA method for 

direct detection of the oyster pathogen Haplosporidium nelsoni. Mar. Biotechnol., 1:147-

54. 

Laird, M. (1961) Microecological factors in oyster epizootics. Can. J. Zool., 39:449-85. 

Landau, H. and P. S. Galtsoff (1951) Distribution of Nematopsis infection on the oyster 

grounds of the Chesapeake Bay and in other waters of the Atlantic and Gulf states. Tex. 

J. Sci., 3:115-30. 

Landsberg, J. H. (1996) Neoplasia and biotoxins in bivalves: is there a connection? J. 

Shellfish Res., 15:203-30. 

Laruelle, F., D. P. Molloy, S. I. Fokin and M. A. Ovcharenko (1999) Histological 

analysis of mantle-cavity ciliates in Dreissena polymorpha: their location, symbiotic 

relationship, and distinguishing morphological characteristics. J. Shellfish Res. 18:251-7. 

48
Lawler, I. F. and J. C. Aldrich (1987) Sublethal effects of bis(tri-N-butyltin)oxide on 

Crassostrea gigas spat. Mar. Pollut. Bull., 6:274-8. 

Lee, R. F. (1993) Passage of xenobiotics and their metabolites from hepatopancreas into 

ovary and oocytes of blue crabs, Callinectes sapidus: possible implications for 

vitellogenesis. Mar. Environ. Res., 35:181-7. 

Lee, M., G. T. Taylor, V. M. Bricelj, S. E. Ford and S. Zahn (1996) Evolution of Vibrio 

spp. and microplankton blooms as causative agents of juvenile oyster disease in 

Crassostrea virginica (Gmelin). J. Shellfish Res., 15:319-29. 

Lichtenfels, J. R., T. K. Sawyer and G. C. Miller (1980) New hosts for larval Sulcascaris

sp. (Nematoda, Anisakidae) and prevalence in the calico scallop (Argopecten gibbus). 

Trans. Amer. Micros. Soc., 99:448-51. 

Little, J. W., S. H. Hopkins and J. G. Mackin (1969) The viability of an echinostome 

metacercaria in the American oyster, Crassostrea virginica. J. Invertebr. Pathol., 

13:455-6. 

Lowe, D. M. and M. N. Moore (1979) The cytology and occurrence of granulocytomas in 

mussels. Mar. Poll. Bull., 10:137-41. 

MacKenzie, K., H. H. Williams, B. Williams, A. H. McVicar and R. Siddall (1995) 

Parasites as indicators of water quality and the potential use of helminth transmission in 

marine pollution studies. Adv. Parasitol., 35:85-114. 

Marigómez, J. A., V. S. Sáez, M. P. Cajaraville and E. Angulo (1990) A planimetric 

study of the mean epithelial thickness (MET) of the molluscan digestive gland over the 

tidal cycle and under environmental stress conditions. Helgol. Meeresunters., 44:81-94. 

Millemann, R. E. (1963) Studies on the taxonomy and life history of echinocephalid 

worms (Nematoda: Spiruroidea) with a complete description of Echinocephalus 

pseudouncinatus Millemann, 1951. J. Parasitol., 49:754-64. 

Mo, C. and B. Neilson (1993) Weight and salinity effects on zinc uptake and 

accumulation for the American oyster (Crassostrea virginica Gmelin). Environ. Pollut., 

82:191-6. 

Morado, J. F., A. K. Sparks and S. K. Reed (1984) A coccidian infection of the kidney of 

the native littleneck clam Protothaca staminea. J. Invertebr. Pathol., 43:207-17. 

Moret, K., K. Williams, C. Couturier and J. Parsons (1999) Newfoundland cultured 

mussel (Mytilus edulis) industry 1997 health survey. Bull. Aquacul. Assoc. Canada, 99-

3:35-7. 

Murchelano, R. A. and S. A. MacLean (1990) Histopathology Atlas of the Registry of 

Marine Pathology. NOAA/NOS/National Ocean Pollution Program Office, Oxford, 

Maryland. 77 pp. 

49
Otto, S. V., J. C. Harshbarger and S. C. Chang (1979) Status of selected unicellular 

eucaryote pathogens, and prevalence and histopathology of inclusions containing obligate 

procaryote parasites, in commercial bivalve mollusks from Maryland estuaries. Haliotis,

8:285-95. 

Peters, E. C. (1988) Recent investigations on the disseminated sarcomas of marine 

bivalve molluscs. In: Disease Processes in Marine Bivalve Molluscs. W. S. Fisher (ed.). 

Am. Fish. Soc. Spec. Publ., 18:74-92. 

Powell, E. N., R. D. Barber, M. C. Kennicutt II and S. E. Ford (1999) Influence of 

parasitism in controlling the health, reproduction and PAH body burden of petroleum 

seep mussels. Deep-Sea Res., I 46:2053-78. 

Powell, E. N., J. M. Klinck and E. E. Hofmann (1996) Modeling diseased oyster 

populations. II. Triggering mechanisms for Perkinsus marinus epizootics. J. Shellfish 

Res., 15:141-65. 

Prytherch, H. F. (1940) The life cycle and morphology of Nematopsis ostrearum, sp. 

nov., a gregarine parasite of the mud crab and oyster. J. Morph., 66:39-65. 

Quick, J. A. and J. G. Mackin (1971) Oyster parasitism by Labyrinthomyxa marina in 

Florida. Fl. Dept. Nat. Resour. Mar. Res. Lab. Prof. Pap, 13:1-55. 

Robledo, J. A. F., M. M. Santarém and A. Figueras (1994) Parasite loads of rafted blue 

mussels (Mytilus galloprovincialis) in Spain with special reference to the copepod, 

Mytilicola intestinalis. Aquaculture, 127:287-302. 

Rózsa, L., J. Reiczigel and G. Majoros (2000) Quantifying parasites in samples of hosts. 

J. Parasitol., 86:228-32. 

Sandoz, M. and S. H. Hopkins (1947) Early life history of the oyster crab, Pinnotheres 

ostreum (Say). Biol. Bull. (Woods Hole) 93:250-8. 

Sindermann, C. J. (1970) Principal Diseases of Marine Fish and Shellfish. Academic 

Press, Inc., New York, NY. 369 pp. 

Sindermann, C. J. (1990) Principal Diseases of Marine Fish and Shellfish. (Second 

Edition) Vol. 2: Diseases of Marine Shellfish. Academic Press, Inc., San Diego, CA. 516 

pp. 

Sparks, A. K. (1985) Synopsis of Invertebrate Pathology Exclusive of Insects. Elsevier 

Science Publishers, Amsterdam, Netherlands. 423 pp. 

Sprague, V. and P. E. Orr (1952) Studies on Nematopsis: III. N. ostrearum and N. 

prytherchi with special reference to host-parasite relation. Natl. Shellfish. Assoc. Conv. 

Add., 1952:26-43. 

50
Stauber, L. A. (1945) Pinnotheres ostreum, parasitic on the American oyster, Ostrea 

(Gryphaea) virginica. Biol. Bull., 88:269-91. 

Stokes, N. A. and E. M. Burreson (2001) Differential diagnosis of mixed Haplosporidium 

costale and Haplosporidium nelsoni infections in the eastern oyster, Crassostrea 

virginica, using DNA probes. J. Shellfish Res., 20:207-13. 

Stunkard, H. W. and J. R. Uzmann (1959) The life-cycle of the digenetic trematode, 

Proctoeces maculatus (Looss, 1901) Odhner, 1911 [syn. P. subtenuis (Linton, 1907) 

Hanson, 1950], and description of Cercaria adranocerca n. sp. Bio. Bull., 116:184-93. 

Teia dos Santos, A. M. and J. Coimbra (1995) Growth and production of raft-cultured 

Mytilus edulis L., in Ria de Aveiro: gonad symbiotic infestation. Aquaculture, 132:195-

211. 

Toews, S., M. Beverley-Burton and T. Lawrimore (1993) Helminth and protist parasites 

of zebra mussels, Dreissena polymorpha (Pallas, 1771), in the Great Lakes region of 

southwestern Ontario, with comments on associated bacteria. Can. J. Zool. 71:1763-6. 

Tripp, M. R. and R. M. Turner (1978) Effects of the trematode Proctoeces maculatus on 

the mussel Mytilus edulis. In: Comparative Pathobiology. Vol. 4 Invertebrate Models for 

Biomedical Research. L. A. Bulla Jr. and T. C. Cheng (eds.). Plenum Press, New York, 

NY. 73-84. 

Villalba, A., S. G. Mourelle, M. J. Carballal and C. Lopez. (1997) Symbionts and 

diseases of farmed mussels Mytilus galloprovincialis throughout the culture process in 

the Ras of Galicia (NW Spain). Dis. Aquat. Org., 31:127-39. 

Warburton, F. E. (1958) Control of the boring sponge on oyster beds. Fish. Res. Board 

Can. Prog. Rpt. Atl. Coast Stns., 69:7-11. 

Weis, P., J. S. Weis, J. Couch, C. Daniels and T. Chen (1995) Pathological and 

genotoxicological observations in oysters (Crassostrea virginica) living on chromated 

copper arsenate (CCA)-treated wood. Mar. Environ. Res., 39:275-8. 

White, M. E., C. L. Kitting and E. N. Powell (1985) Aspects of reproduction, larval 

development and morphometrics in the pyramidellid Boonea (=Odostomia) impressa

(Gastropoda: Opisthobranchia). Veliger, 28:37-51. 

White, M. E., E. N. Powell, E. A. Wilson and S. M. Ray (1989) The spatial distribution 

of Perkinsus marinus, a protozoan parasite, in relation to its oyster host (Crassostrea 

virginica) and an ectoparasitic gastropod (Boonea impressa). J. Mar. Biol. Assoc. UK, 

69:703-17. 

Wilson-Ormond, E.A., M.S. Ellis, E.N. Powell, Y. Kim and S. Li (2000) Effects of gasproducing platforms on continental shelf megafauna in the northwest Gulf of Mexico: 

reproductive status and health. Internat. Rev. Hydrobiol., 85:293-323. 

51
Winstead, J. T. (1995) Digestive tubule atrophy in eastern oysters, Crassostrea virginica 

(Gmelin, 1791), exposed to salinity and starvation stress. J. Shellfish Res., 14:105-11. 

Winstead, J. T. and J. A. Couch (1981) Proctoeces sp. (Trematoda: Digenea) in the 

American oyster, Crassostrea virginica. Trans. Am. Micros. Soc., 100:296-305. 

Winstead, J. T. and J. A. Couch (1988) Enhancement of protozoan pathogen Perkinsus 

marinus infections in American oysters Crassostrea virginica exposed to the chemical 

carcinogen n-nitrosodiethylamine (DENA). Dis. Aquat. Org., 5:205-13. 

Winstead, J. T., R. M. Overstreet and L. A. Courtney (1998) Novel gonadal parasites in 

the eastern oyster Crassostrea virginica from two Gulf of Mexico bays. J. Shellfish Res., 

17:341-342. 

Wolf, P. H., J. T. Winstead and J. A. Couch (1987) Proctoeces sp. (Trematoda: Digenea) 

in Australian oysters, Saccostrea commercialis and Crassostrea amasa. Trans. Am. 

Micros. Soc., 106:379-80. 

Yoo, M. S. and T. Kajihara (1985) Effect of the pea crab (Pinnotheres pholadis) on the 

reproductive capacity of the blue mussel (Mytilus edulis galloprovincialis). Bull. Korean 

Fish. Soc., 18:581-5. 

Zander, C. D. (1998) Ecology of host parasite relationships in the Baltic Sea. Naturwiss., 

85:426-36. 

Zaroogian, G., P. Yevich and S. Anderson (1993) Effect of selected inhibitors on

cadmium, nickel, and benzo[a]pyrene uptake into brown cells of Mercenaria mercenaria. 

Mar. Environ. Res., 35:41-5. 

52
CHAPTER 3. PERKINSUS MARINUS ASSAY 

K.A. Ashton-Alcox, Y. Kim, and E. N. Powell 

Haskin Shellfish Research Laboratory 

Rutgers University 

Port Norris, NJ 

ABSTRACT 

This chapter describes the procedures followed for the assay of Perkinsus marinus 

(etiological agent of Dermo disease) presence and infection intensity in oysters collected 

on the East and Gulf Coasts of the US and in Puerto Rico for NOAA’s National Status 

and Trends Mussel Watch Project. Analyses are conducted on mantle tissue excised from 

the oyster and placed in Ray’s Fluid Thioglycollate Medium (RFTM) to enlarge P. 

marinus present in the tissue. The enlarged cells are stained by the addition of Lugol’s 

iodine solution. The tissue is examined microscopically and rated on a semi-quantitative 

scale. 

1.0 INTRODUCTION 

Perkinsus marinus is the most widespread pathogen of East and Gulf coast oysters and is 

the etiologic agent of Dermo disease [see Ford and Tripp (1996) for review]. Once 

considered a form of fungus (Mackin et al,. 1950), this protozoan parasite is now 

generally considered to be a member of the Apicomplexa (Levine, 1978). However, 

Reece et al. (1997) have recently emphasized the parasite’s dinoflagellate affinities so 

that its phylogenetic placement remains controversial. Dermo is transmitted from oyster 

to oyster (Perkins and Menzel, 1966) and causes significant mortalities in oyster 

populations. Powell et al. (1992) and Kim et al. (1999) have considered the spatial 

distribution and temporal trend in P. marinus prevalence and infection intensity at Mussel 

Watch sites. Recent research continues to support the likely relationship of Dermo 

disease to environmental stress (Powell et al., 1996; Lenihan et al., 1999), including 

contaminant exposure (Chu, 1999; Fisher et al., 1999; Chu and Hale, 1994; Wilson et al., 

1990; Winstead and Couch, 1988; Scott et al., 1985). Most recently, Tall et al. (1999) 

have suggested a possible relationship between Dermo disease and Vibrio vulnificus, an 

important source of infection in susceptible humans eating raw oysters (Shapiro et al., 

1998). 

The determination of the health of oyster populations requires an analysis of P. marinus 

prevalence and infection intensity. The standard method for monitoring purposes is the 

one developed by Ray (1966). Briefly, a section of mantle tissue is incubated in Ray’s 

Fluid Thioglycollate Medium (RFTM) for 7 days under anaerobic, dark conditions. In the 

RFTM, the P. marinus cells enlarge to form thick-walled hypnospores but do not 

replicate (Ray, 1954; Stein and Mackin 1957). The hypnospores are stained with Lugol’s 

iodine and a semi-quantitative (Craig et al., 1989) assessment is made microscopically. A 

53
quantitative estimate can be derived from the semi-quantitative designation using the 

formula found in Choi et al. (1989). 

2.0 EQUIPMENT AND SUPPLIES 

2.1. REAGENTS 

2.1.1. CHEMICALS

Denatured alcohol (for burner and sterilization), A962P-4. Fisher Scientific, Pittsburgh, 

PA. 

Fluid thioglycollate medium (FTM), powder, T9032. Sigma Chemical Company, St. 

Louis, MO. 

Iodine, (I2) [7553-56-2], I3380, purity 99%. Sigma Chemical Company, St. Louis, MO. 

Penicillin (C16H17N2O4SNa) [69-57-8], P3032. Sigma Chemical Company, St. Louis, 

MO. 

Potassium iodide (KI) [7681-11-0], P8256. Sigma Chemical Company, St. Louis, MO. 

Sodium chloride (NaCl) [7647-14-5] S-271. Fisher Scientific, Pittsburgh, PA. 

Streptomycin sulfate (C21H39N7O12)2-(H2SO4)3 [3810-74-0] BP910-50. Fisher Biotech, 

Pittsburgh, PA. 

2.1.2. SOLUTIONS

2.1.2.1. Ray’s (1952) Thioglycollate medium preparation 

A mixture of 22 g NaCl, 29.3 g dehydrated Fluid Thioglycollate Medium (FTM), and 1 L 

distilled water is heated while stirring until the medium dissolves and the solution boils 

and becomes a transparent golden-yellow color. After cooling, the solution is dispensed, 

5 mL at a time, into 15-mL culture tubes which are subsequently autoclaved and sealed. 

They are kept in the dark until use. Unused, autoclaved tubes of RFTM can be stored for 

many months in the dark without deterioration. Discard them if they become cloudy or 

the RFTM congeals. 

RFTM maintains anaerobic conditions in the culture tube as well as providing needed 

nutrients and an appropriate osmotic environment. Therefore, tubes are sealed tightly and 

opened only briefly for addition of antibiotic and tissue as described below. After tissue 

is added, tubes are sealed and returned immediately to the dark for tissue incubation. 

2.1.2.2. Antibiotic solution 

A 0.33 g quantity of Streptomycin sulfate and 0.159 g of Penicillin G are added to 500 

mL of sterilized, deionized water and shaken lightly until the powder is dissolved. The 

54
solution is kept refrigerated until use. The solution can be stored safely for several 

months if refrigerated. 

2.1.2.3. Lugol’s iodine solution 

Four grams of potassium iodide and 2 g of iodine crystals are dissolved in 100 mL of 

distilled water, allowed to stand for 24 h, and filtered. The solution is stored in a dark 

bottle at room temperature to avoid particle precipitation. The solution remains stable for 

many weeks but should be filtered occasionally to remove particles that may precipitate. 

These particles may be confused with P. marinus hypnospores by less experienced slide 

readers although they are disks, not spheres; they are always black, never blue; and they 

are birefringent (Bushek et al., 1994). 

2.2. EQUIPMENT 

Alcohol lamp 

Autoclave 

Automatic repipet, Barnstead/Thermolyne Repipet II (9010), 13-687-62B. Fisher 

Scientific, Pittsburgh, PA. 

Coverslips, various sizes, eg. 12-545-B. Fisher Scientific, Pittsburgh, PA. 

Culture tubes with Screw Cap, Pyrex, 14-932A. Fisher Scientific, Pittsburgh, PA. 

Dissecting tools- scissors, scalpels, probes, forceps 

Heater/stirrer, 11-500-7SH. Fisher Scientific, Pittsburgh, PA. 

Microscope, Carl Zeiss Standard KF2, 12-070-273. Fisher Scientific, Pittsburgh, PA. 

Microscope slides, Fisherbrand Frosted, 12-550-34. Fisher Scientific, Pittsburgh, PA. 

Oyster knives Dexter/Russell “Sani-Safe” S134. Memphis Net & Twine Co., Inc. 

Memphis, TN. 

Pasteur pipets, 13-678-20A. Fisher Scientific, Pittsburgh, PA. 

Repipet, 1 mL capacity, adjustable volume, P-1000. Rainin Instrument Co., Woburn, 

MA. 

Repipet tips, Redi-Tip Universal, 21-197-8F. Fisher Scientific, Pittsburgh, PA. 

Vortex mixer, Vortex-Genie 2 Model G-560, 12-812. Fisher Scientific, Pittsburgh, PA. 

55
3.0 TISSUE COLLECTION 

Twelve oysters are randomly selected from each group of 20-25 collected per site. The 

anterior-posterior length (Morales-Alamo and Mann, 1989) of each oyster is measured 

with a ruler. Each oyster is opened with an oyster knife by cutting the adductor muscle at 

its connection with the upper (right) shell. A gross examination of each oyster is made 

and each oyster is given a numerical condition code according to the system described by 

Quick and Mackin (1971) (Table 3-1). Using sterile dissecting scissors and forceps, a 5 x 

5-mm piece of mantle-edge tissue is excised from just over the palps. Sterile instruments 

must be used when going from oyster to oyster to avoid cross-contamination. An alcohol 

dip and flaming is sufficient. The tissue is placed in a culture tube containing 5 mL 

RFTM to which 0.5 mL of the Penicillin-Streptomycin solution has been added. The tube 

is recapped and inverted so that the tissue is submerged in the RFTM. When processing 

many samples, it is convenient to add the antibiotic solution to the vials prior to, but not 

earlier than 1 h before, opening the oysters. Of the original 12 oysters, 5 are randomly 

chosen and further processed for gonadal analysis and histopathology (see Kim et al., this 

volume). The culture tubes are placed in the dark at room temperature and incubated for 

at least 5 days. If the tissue has not been analyzed by the end of 7 days, the tube is placed 

in a refrigerator in the dark. Tissues ready to be analyzed can be kept for at least 3 

months without deterioration if the culture tubes are kept dark and refrigerated. 

4.0 TISSUE ANALYSIS 

After the incubation period, the oyster tissue is removed from the RFTM using a sterile 

probe and placed on a microscope slide. The tissue sample is teased apart using sterile 

probes to assure even staining with Lugol’s iodine solution. One or two drops of Lugol’s 

solution is added to the tissue with a Pasteur pipette, the tissue is covered with a cover 

slip, and then examined microscopically. (Note: Microscope slides are thoroughly 

cleaned and re-used although coverslips are disposed of after use) 

Perkinsus marinus hypnospores appear as blue/black spheres 5 to 300 μm in diameter 

when viewed through a microscope at 40 to 100x magnification (Fig. 3-1 and Bushek et 

al., 1994). An infection intensity is assigned to each sample based on the number or 

coverage of enlarged P. marinus observed in the tissue using the scale in Table 3-2. For 

example, in Figure 3-1: Photo 1 would be rated VL (.33) or L- (.67); Photo 2 would be 

rated L+ (1.33); Photo 3 would be rated M- (2.67); Photo 4 would be rated MH (4.00); 

Photo 5 would be rated H (5.00). 

To maintain quality control, blind assays may be conducted among slide readers to 

correct for the technician bias that may be present with any semi-quantitative technique. 

Other laboratories are encouraged to standardize their analyses with laboratories already 

using the technique so that data are comparable (e.g., Fisher and Oliver, 1996). 

56
Table 3-1. Oyster condition rating key (from Quick and Mackin, 1971). 

Condition 

index 

Code 

no. 

Oyster appearance 

Very good 1 Animal firm and filling shell cavity; coloration creamy white 

and evenly textured; usually ready to spawn 

Good 2 Not quite as firm or large as above; usually ready to spawn 

Good minus 3 Coloration less opaque, often slightly yellow or gray 

 

Fair plus 4 Animal distinctly not filling shell cavity; coloration often 

mottled, with blood vessels and muscle fibers showing 

through the more translucent epithelium 

Fair 5 Oyster well-developed but not opaque or tending toward 

white; grayish and translucent; flesh flaccid 

Fair minus 6 Translucency more pronounced 

Poor plus 7 Oyster not well-developed, darker gray, often greenish; 

pericardial cavity clear; small portion of shell cavity filled 

Poor 8 Negative qualities more accentuated 

Very poor 9 Animal distinctly atrophied; coloration dark and uneven, 

very translucent; seldom more than third of shell cavity 

occupied; adductor muscle often discolored and transparent 

even in the normally white sector 

57
Figure 3-1. Enlarged and stained Perkinsus marinus hypnospores. Hypnospores appear as 

black dots in the photographs. Scale bars = 3 mm (courtesy of S. E. Ford, Haskin 

Shellfish Research Laboratory, Rutgers University, NJ) 

58
Table 3-2. Semi-quantitative scale of infection intensity for Perkinsus marinus [adapted 

from Mackin (1962) by Craig et al. (1989)]. 

Letter Infection Numerical Description 

designation intensity value 

N Negative 0.00 No hypnospores present 

VL Very light 0.33 1-10 hypnospores 

L- 0.67 11-74 hypnospores 

L Light 1.00 75-125 hypnospores 

L+ 1.33 >125 hypnospores but much less than 25% 

of tissue is hypnospores 

LM- 1.67 <25% of tissue is hypnospores 

LM Light/moderate 2.00 25% of tissue is hypnospores 

LM+ 2.33 >25% but much less than 50% of tissue is 

hypnospores 

M- 2.67 >25% but <50% of tissue is hypnospores 

M Moderate 3.00 50% of tissue is hypnospores 

M+ 3.33 >50% but much less than 75% of tissue is 

hypnospores 

MH- 3.67 >50% but <75% of tissue is hypnospores 

MH Moderately heavy 4.00 75% of tissue is hypnospores 

MH+ 4.33 >75% but much less than 100% of tissue is 

hypnospores 

H- Heavy 4.67 >75% of tissue is hypnospores but some 

oyster tissue is still visible 

H 5.00 Nearly 100% of tissue is hypnospores 

Alternative techniques have been developed for the assay of P. marinus prevalence and 

infection intensity in addition to Ray’s (1966) tissue technique. Gauthier and Fisher 

(1990) used an adaptation of Ray’s method to non-destructively follow the progression of 

P. marinus infection by sampling oyster hemolymph. Choi et al. (1989) and Fisher and 

Oliver (1996) describe quantification of the RFTM method by using a total body burden 

technique to assess total numbers of P. marinus cells present in an oyster. Whole body 

counts minimize false negatives that commonly occur at low P. marinus infection levels 

(Nickens et al., 2002; Choi et al., 1989). The accuracy and precision of the three RFTMbased methods is discussed in Bushek et al. (1994) and Oliver et al. (1998). Although the 

most accurate method for estimation of P. marinus infection levels in individual oysters 

59
is total body burden, the removal of a small piece of mantle tissue is rapid, inexpensive, 

and can be done with little tissue loss from animals destined for other analyses. This 

method accurately reflects the prevalence and infection intensity in an oyster population 

(Bushek et al., 1994; Oliver et al., 1998). Recently described molecular techniques are 

not yet in general use but promise greater precision and accuracy (eg., Ottinger, et al., 

2001) although time and expense, again, may be prohibitive for monitoring purposes. 

Although the RFTM method is recommended for survey and monitoring work (Bushek et 

al., 1994; Oliver et al., 1998) and is widely used, some cautionary comments are 

necessary. 

1. The relatively small size of the sample used in this method may lead to false 

negatives at less than 1000 P. marinus per gram wet weight of oyster tissue (Bushek 

et al., 1994; Choi et al., 1989; see also Ottinger et al., 2001) or inaccurate sample 

ranking due to the non-uniform distribution of P. marinus within (Choi et al., 1989) 

and between (Oliver et al., 1998) oyster tissues. The use of the Ray (1966) FTM 

method has been shown, however, to accurately estimate the relative overall intensity 

of P. marinus in a sample, if not necessarily in individual oysters (Bushek et al., 

1994; Oliver et al., 1998), and is considered to be adequate for survey and monitoring 

work. 

2. All microscopically identifiable stages of P. marinus present in the oyster appear to 

enlarge in RFTM with minimal reproduction so that the number of cells present after 

incubation accurately represents the number of cells in the oyster tissue (Ray, 1954; 

Stein and Mackin, 1957). However, the mean diameter of hypnspores can vary from 5 

μ to 300 μ (Fig. 3-1, see also Fig. 2 Bushek et al., 1994; Ray, 1954). The degree of 

enlargement appears to vary seasonally and may be inversely related to infection 

intensity (Bushek et al., 1994). The size of the hypnospores is not taken into account 

by the slide reader and, because the ranking scale is semi-quantitative, the numbers of 

cells are not counted except at the “Light” and lower levels of infecton. This means 

that a tissue sample with fewer large hypnospores may have the same numerical rank 

as a sample with many smaller cells. In cases where this source of error is of concern, 

calibration using tissue weights can be conducted (Choi et al., 1989): 

Hypnospores (g wet wt.)-1= 1409.9 (100.64296x) 

 where x is the semi-quantitative numerical value from Table 3-2 (Choi et al., 1989). 

5.0 DEFINITIONS AND CALCULATIONS 

Three calculations of parasite distribution are generally used to describe Dermo disease in 

oyster populations: infection intensity, prevalence, and weighted prevalence (also termed 

mean abundance-Bush et al., 1997; Rosza et al., 2000). Infection intensity for each site 

sampled is calculated as the sum of the infection rankings from Table 3-2, divided by the 

number of oysters with infections. Prevalence describes the proportion of individuals in 

the sample that are infected and is calculated as the number of oysters infected by P. 

marinus divided by the total number of oysters analyzed. Weighted prevalence or mean 

abundance is the infection intensity multiplied by the prevalence. This gives a measure of 

60
the relative severity of P. marinus infection in a population. Due to the truncated nature 

of the semi-quantitative scale in Table 3-2 and because infection intensity in a sampled 

population may not be normally distributed, the calculation of the median infection is 

often more desirable than the mean infection. Confidence intervals for the median can 

normally be calculated by bootstrapping (Efron and Tibshirani, 1986). 

6.0 CONCLUSIONS 

The described technique provides a semi-quantitative method to determine the presence 

and infection intensity of P. marinus in oysters. Despite its drawbacks, the Ray (1966) 

FTM method is currently the accepted method to determine population health in routine 

monitoring. This method is considerably more accurate than examination of tissue 

sections in routine histopathological analysis (Ray, 1954) and less expensive and timeconsuming than the more accurate body burden method (Choi et al., 1989; Bushek et al., 

1994; Oliver et al., 1998). 

7.0 REFERENCES

Bush, A. O., K. D. Lafferty, J. M. Lotz and A. W. Shostak (1997) Parasitology meets 

ecology on its own terms: Margolis et al. revisited. J. Parasitol., 83:575-583. 

Bushek, D., S. E. Ford and S. K. Allen, Jr. (1994) Evaluation of methods using Ray’s 

fluid thioglycollate medium for diagnosis of Perkinsus marinus infection in the Eastern 

oyster, Crassostrea virginica. Ann. Rev. Fish Dis. 4:201-217. 

Choi, K-S., E. A. Wilson, D. H. Lewis, E. N. Powell and S. M. Ray (1989) The energetic 

cost of Perkinsus marinus parasitism in oysters: quantification of the thioglycollate 

method. J. Shellfish Res., 8: 125-31. 

Chu, F.-L. E. (1999) Environmental factors and the infectious disease caused by the 

protozoan parasite, Perkinsus marinus, in Eastern oysters (Crassostrea virginica). Bull. 

Eur. Assoc. Fish. Pathol., 19:265-268.

Chu, F.-L. E. and R. C. Hale (1994) Relationship between pollution and susceptibility to 

infectious disease in the Eastern oyster, Crassostrea virginica. Mar. Environ. Res., 

38:243-256. 

Craig, M. A., E. N. Powell, R. R. Fay and J. M. Brooks (1989) Distribution of Perkinsus 

marinus in Gulf coast oyster populations. Estuaries, 12: 82-91. 

Efron, B. and R. Tibshirani (1986) Bootstrap methods for standard errors, confidence 

intervals, and the measures of statistical accuracy. Stat. Sci., 1:54-77. 

Fisher, W. S. and L. M. Oliver (1996) A whole-oyster procedure for diagnosis of 

Perkinsus marinus disease using Ray’s fluid thioglycollate culture medium. J. Shellfish 

Res., 15:109-117. 

61
Fisher, W. S., L. M. Oliver, W. W. Walker. C. S. Manning and T. F. Lytle (1999) 

Decreased resistance of eastern oysters (Crassostrea virginica) to a protozoan pathogen 

(Perkinsus marinus) after sublethal exposure to tributyltin oxide. Mar. Environ Res., 

47:185-201. 

Ford, S. E. and M. R. Tripp, (1996) Diseases and Defense Mechanisms. In: The Eastern 

Oyster: Crassostrea virginica. R. I. E. Newell and A. F. Eble, (eds.), Maryland Sea Grant 

College, College Park, MD, p. 581-660. 

Gauthier, J. D. and W. S. Fisher (1990) Hemolymph assay for diagnosis of Perkinsus 

marinus in oysters Crassostrea virginica (Gmelin, 1791). J. Shellfish Res., 9:367-372. 

Kim, Y., E. N. Powell, T. L. Wade, B. J. Presley and J. M. Brooks (1999) Influence of 

climate change on interannual variation in contaminant body burden in Gulf of Mexico 

oysters. Mar. Environ. Res., 48:459-488. 

Lenihan, H. S., F. Micheli, S. W. Shelton and C. H. Peterson (1999) The influence of 

multiple environmental stressors on susceptibility to parasites: an experimental 

determination with oysters. Limnol. Oceanogr., 44:910-924. 

Levine, N. D. (1978) Perkinsus gen. n. and other new taxa in the protozoan phylum 

Apicomplexa. J. Parasitol. 64:549. 

Mackin, J. G., H. M. Owen and A. Collier (1950) Preliminary note on the occurrence of a 

new protistan parasite, Dermocystidium marinum n. sp., in Crassostrea virginica

(Gmelin). Science, 111:328-329. 

Mackin, J. G. (1962) Oyster disease caused by Dermocystidium marinus and other 

microorganisms in Louisiana. Publ. Inst. Mar. Sci. Univ. Texas, 7:132-229. 

Morales-Alamo, R. and R. Mann (1989) Anatomical features in histological sections of 

Crassostrea virginica (Gmelin, 1791) as an aid in measurements of gonad area for 

reproductive assessment. J. Shellfish Res. 8:71-82. 

Nickens, A. D., J. F. LaPeyre, E. S. Wagner and T. R. Tiersch (2002) An improved 

procedure to count Perkinsus marinus in Eastern oyster hemolymph. J. Shellfish Res., 

21:725-732. 

Oliver, L.M., W. S. Fisher, S. E. Ford, L. M. Ragone Calvo, E. M. Burreson, E. B. Sutton 

and J. Gandy (1998) Perkinsus marinus tissue distribution and seasonal variation in 

oysters Crassostrea virginica from Florida, Virginia, and New York. Dis. Aq. Org.. 

34:51-61. 

Ottinger, C. A., T. D. Lewis, D. A. Shapiro, M. Faisal and S. L. Kaattari (2001) Detection 

of Perkinsus marinus extracellular proteins in tissues of the Eastern oyster, Crassostrea 

virginica: potential use in diagnostic assays. J. Aquat. Animal Heath, 13:133-141. 

62
Perkins, F. O. and R. W. Menzel, (1966) Morphological and cultural studies of a motile 

stage in the life cycle of Dermocystidium marinum. Proc. Natl. Shelllfish Asssoc. 56:23-

30. 

Powell, E. N., J. D. Gauthier, E. A. Wilson, A. Nelson, R. R. Fay and J. M. Brooks 

(1992) Oyster disease and climate change. Are yearly changes in Perkinsus marinus

parasitism in oysters (Crassostrea virginica) controlled by climatic cycles in the Gulf of 

Mexico? Mar. Ecol., 13:243-270. 

Powell, E. N., J. M. Klinck and E. E. Hofmann (1996) Modeling diseased oyster 

populations. II. Triggering mechanisms for Perkinsus marinus epizootics. J. Shellfish 

Res., 15:141-165. 

Quick, J. A. and J. G. Mackin (1971) Oyster parasitism by Labyrinthomyxa marina in 

Florida. Fl. Dept. Nat. Resour. Mar. Res. Lab. Prof. Pap. Ser., 13:1-55. 

Ray, S. M. (1952) A culture technique for the diagnosis of infections with 

Dermocystidium marinum, Mackin, Owen and Collier, in oysters. Science, 166:360-361. 

Ray, S. M. (1954) Biological studies of Dermocystidium marinus, a fungus parasite of 

oysters. Rice Inst. Pamph. Monogr. Biol. Special Issue, Rice University, Houston, TX, 

114 pp. 

Ray, S. M. (1966) A review of the culture method for detecting Dermocystidium marinus, 

with suggested modifications and precautions. Proc. Natl. Shellfish. Assoc., 54: 55-69. 

Reece, K. S., M. E. Siddall, E. M. Burreson and J. E. Graves (1997) Phylogenetic 

analysis of Perkinsus based on actin gene sequences. J. Parasitol. 83:417-423. 

Rózsa, L., J. Reiczigel and G. Majoros (2000) Quantifying parasites in samples of hosts. 

J. Parasitol., 86:228-32. 

Scott, G. I., D. P. Middaugh and T. I. Sammons (1985) Interactions of chlorine-produced 

oxidants (CPO) and salinity in affecting lethal and sublethal effects in the eastern oyster, 

Crassostrea virginica (Gmelin) infected with the protistan parasite Perkinsus marinus. 

In: Marine Pollution and Physiology, Recent Advances, F. J. Vernberg, C. P. Thurnberg, 

A. Calabrese, and W. B. Vernberg (eds.). University of South Carolina Press, Columbia, 

SC. 

Shapiro, R., L. Alketruse, R. Hutwagner, R. Bishop, R. Hammond, S. Wilson, B. Ray, S. 

Thompson, R. Tauxe, P. Griffin and the Vibrio Working Group (1998) The role of Gulf 

coast oysters harvested in warmer months in Vibrio vulnificus infection in the United 

States, 1988-1996. J. Infect. Dis., 178:752-759. 

Stein J. E. and J. G. Mackin (1957) An evaluation of the culture method used in 

determining the intensity of Dermocystidium marinum in the oyster C. virginica. Texas 

A&M Research Foundation, Project 23, Tech Report No. 22. 5 pp. 

63
Tall, B.D., J. F. LaPeyre, J. W. Bier, M. D. Miliotis, D. E. Hanes, M. H. Kothary, D. B. 

Shah and M. Faisal (1999) Perkinsus marinus extracellular protease modulates survival 

of Vibrio vulnificus in Eastern oyster (Crassostrea virginica) hemocytes. Appl. Environ. 

Microbiol., 65:4261-4263. 

Wilson, E. A.. E. N. Powell, M. A Craig, T. L. Wade and J. M. Brooks (1990) The 

distribution of Perkinsus marinus in Gulf coast oysters: its relationship with temperature, 

reproduction, and pollutant body burden. Int. Rev. Gesamten Hydrobiol., 75:533-550. 

Winstead, J. T. and J. A. Couch (1988) Enhancement of protozoan pathogen Perkinsus 

marinus in oysters Crassostrea virginica exposed to the chemical carcinogen nnitrosodiethylamine (DENA). Dis. Aquat. Org., 5:205-213.

64

United States Department of Commerce 

Carlos M. Gutierrez 

 Secretary 

National Oceanic and Atmospheric Administration 

Vice Admiral Conrad C. Lautenbacher, Jr. USN (Ret.) 

Under Secretary of Commerce for Oceans and Atmospheres 

National Ocean Service 

Jack Dunnigan 

Assistant Administrator for Ocean Service and Coastal Zone Management