basic anatomy of an oyster

108

Basic Anatomy of an Oyster

Basic anatomy of an oyster.

rectum�

pericardial �cavity�

anus�

denticle�

stomach�

digestive gland�

mouth�

mantle�

gill�adductor muscle�

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SECTION 3 - MOLLUSCAN DISEASES

Basic Anatomy of an OysterSECTION 3 - MOLLUSCAN DISEASESM.1 GENERAL TECHNIQUESM.1.1 Gross ObservationsM.1.1.1 BehaviourM.1.1.2 Shell Surface ObservationsM.1.1.3 Inner Shell ObservationsM.1.1.4 Soft-Tissue SurfacesM.1.2 Environmental ParametersM.1.3 General ProceduresM.1.3.1 Pre-Collection PreparationM.1.3.2 Background InformationM.1.3.3 Sample Collection for Health SurveillanceM.1.3.4 Sample Collection for Disease DiagnosisM.1.3.5 Live Specimen Collection for ShippingM.1.3.6 Preservation of Tissue SamplesM.1.3.7 Shipping Preserved SamplesM.1.4 Record KeepingM.1.4.1 Gross ObservationsM.1.4.2 Environmental ObservationsM.1.4.3 Stocking RecordsM.1.5 References

DISEASES OF MOLLUSCSM.2 Bonamiosis (Bonamia sp., B. ostreae)M.3 Marteiliosis (Marteilia refringens, M. sydneyi)M.4 Mikrocytosis (Mikrocytos mackini, M. roughleyi)M.5 Perkinsosis (Perkinsus marinus, P. olseni)M.6 Haplosporidiosis (Haplosporidium costale,

H. nelsoni)M.7 Marteilioidosis (Marteilioides chungmuensis, M.

branchialis)M.8 Iridovirosis (Oyster Velar Virus Disease)

ANNEXESM.AI OIE Reference Laboratories for

Molluscan DiseasesM.AII List of Regional Resource Experts for Molluscan

Diseases in Asia-PacificM.AIII List of Useful Diagnostic Guides/Manuals to

Molluscan Health

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110111111111111114114116116116116116116117118118118119119119

121125129133138

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M.1 GENERAL TECHNIQUES

(SE McGladdery)

Fig.M.1.1.1. Gaping hard shell clam,Mercenaria mercenaria, despite air exposureand mechnical tapping.

(MG Bondad-Reantaso)

Fig.M.1.1.2a. Mollusc encrustment (arrows) ofwinged oyster, Pteria penguin, Guian PearlFarm, Eastern Samar, Philippines (1996).

(D Ladra)

Fig.M.1.1.2b. Pteria penguin cultured at GuianPearl Farm, Eastern Samar, Philippines with ex-tensive shell damage due to clionid (boring)sponge (1992).

(MG Bondad- Reantaso)

Fig.M.1.1.2c,d. Pteria penguin shell with densemulti-taxa fouling, Guian Pearl Farm, EasternPhilippines (1996).

(SE McGladdery)

Fig.M.1.1.2e. Polydora sp. tunnels and shelldamage at hinge of American oyster,Crassostrea virginica, plus barnacle encrustingof other shell surfaces.


Fig.M.1.1.2f. Winged oyster, Pteria penguin,shell with clionid sponge damage. Guian PearlFarm, Eastern, Philippines (1996).

c

d

>

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M.1 General Techniques

General molluscan health advice and othervaluable information are available from the OIEReference Laboratories, Regional ResourceExperts in the Asia-Pacific, FAO and NACA.A list is provided in Annexes M.A1 and M.AII,and up-to-date contact information may beobtained from the NACA Secretariat inBangkok (e-mail: [email protected]). Otheruseful guides to diagnostic procedures whichprovide valuable references for molluscan dis-eases are listed in Annex M.AIII.

M.1.1 Gross Observations

M.1.1.1 Behaviour (Level I)

It is difficult to observe behavioural changes inmolluscs in open-water, however, close atten-tion can be made of behaviour of both broodstockand larvae in hatcheries. Since disease situa-tions can erupt very quickly under hatchery con-ditions, regular and close monitoring is worthLevel I efforts (see Iridovirus - M.8).

Feeding behaviour of larval molluscs is also agood indicator of general health. Food accumu-lation in larval tanks should be noted and samplesof larvae examined, live, under a dissecting mi-croscope for saprobiotic fungi and protists (e.g.ciliates) and/or bacterial swarms. Pre-settlementstages may settle to the bottom prematurely orshow passive circulation with the water flow cur-rents in the holding tanks.

Juvenile and adult molluscs may also cease feed-ing, and this should be cause for concern undernormal holding conditions. If feeding does notresume and molluscs show signs of weakening(days to weeks depending on water temperature)samples should be collected for laboratory ex-amination. Signs of weakening include gaping (i.e.bivalve shells do not close when the mollusc istouched or removed from the water) (Fig.M.1.1.1), accumulation of sand and debris inthe mantle and on the gills, mantle retractionaway from the edge of the shell, and decreasedmovement in mobile species (e.g. scallop swim-ming, clam burrowing, abalone grazing, etc.).

Open-water mortalities that assume levels ofconcern to the grower should be monitored todetermine if there are any patterns to the losses.Sporadic moralities following periods of intensehandling should be monitored with minimal addi-tional handling if at all possible. If the mortalitiespersist, or increase, samples should be collectedfor laboratory analysis. Mortalities that appear tohave a uniform distribution should be examined

immediately and environmental factors pre- andpost-mortality recorded. Mortalities that appearto spread from one area to another suggest thepresence of an infectious disease agent andshould be sampled immediately. Affected animalsshould be kept as far away as possible from un-affected animals until the cause of the mortali-ties can be determined.

M.1.1.2 Shell Surface Observations(Level I)

Fouling organisms (barnacles, limpets, sponges,polychaete worms, bivalve larvae, tunicates,bryozoans, etc.) are common colonists of mol-lusc shell surfaces and do not normally presenta threat to the health of the mollusc (Fig.M.1.1.2a,b). Suspension and shallow water cul-ture, however, can increase exposure to foulingand shells may become covered by other ani-mals and plants (Fig. M.1.1.2c,d). This can af-fect health directly by impeding shell opening andclosing (smothering) or indirectly through com-petition for food resources. Both circumstancescan weaken the mollusc so cleaning may be re-quired. Such defouling should be undertaken asrapidly as possible, to minimise the period of re-moval from the water, during cooler periods ofthe day. Rapid cleaning is usually achieved us-ing high pressure water or mechanical scapers.Defouled molluscs should be returned to cleanwater. Fouling organisms should not be discardedin the same area as the molluscs, since this willaccelerate recolonisation. Signs of weakeningthat persist or increase after cleaning, should beinvestigated further by laboratory examination.

Shell damage by boring organisms, such assponges and polychaete worms (Fig. M.1.1.2e,f) is normal in open-water growing conditions.Although usually benign, under certain conditions(especially in older molluscs) shells may be ren-dered brittle or even become perforated. Suchdamage can weaken the mollusc and render itsusceptible to pathogen infections.

Shell deformities (shape, holes in the surface),fragility, breakage or repair should be noted, butare not usually indicative of a disease condi-tion (Fig.M.1.1.2g, h). Abnormal colouration andsmell, however, may indicate a possible soft-tis-sue infection which may require laboratory ex-amination.

M.1.1.3 Inner Shell Observations (Level I)

The presence of fouling organisms (barnacles,sponges, polychaete worms, etc.) on the innershell surface is a clear indication of a weak/

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Fig.M.1.1.2g,h. Pinctada maxima, shell withclionid sponge damage due to excavation oftunnels exhalent-inhalent openings (holes) tothe surface (arrows). Other holes (small arrows)are also present that may have been causedby polychaetes, gastropod molluscs or otherfouling organisms. Guian Pearl Farm, EasternPhilippines (1996).


Fig.M.1.1.3a. Winged oyster, Pteria penguin,shell showing clionid sponge damage throughto the inner shell surfaces, Guian Pearl Farm,Eastern Philippines (1996).

(B Jones)

Fig.M.1.1.3a1. Abalone (Haliotis roei) from abatch killed by polydoriid worms.

(D Ladra)

Fig.M.1.1.3b,c. b. Shells of Pinctada maximashowing a erosion of the nacreous inner sur-faces (arrows), probably related to chronicmantle retraction; c. Inner surface of shellshowing complete penetration by boringsponges (thin arrows).

b

c

g

h

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Fig.M.1.1.3d,e,f. Pinctada maxima (d), Pteriapenguin (e) and edible oyster (Crassostrea sp.)(f) shells showing Polydora-related tunnel dam-age that has led to the formation of mud-filledblisters.


Fig.M.1.1.3g. Inner shell of winged pearl oys-ter showing: tunnels at edge of the shell (straightthick arrow); light sponge tunnel excavation(transparent arrow); and blisters (small thickarrow) at the adductor muscle attachment site.Guian Pearl Farm, Eastern Philippines (1996).

(SE McGladdery)

Fig.M.1.1.3.h. Extensive shell penetration bypolychaetes and sponges causing weakeningand retraction of soft-tissues away from theshell margin of an American oyster Crassostreavirginica.

d

e

f

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sick mollusc (Fig.M.1.1.3a and Fig.M.1.1.3a1).The inner surfaces are usually kept cleanthrough mantle and gill action. Perforation ofthe inner surface can be sealed off by deposi-tion of additional conchiolin and nacre(Fig.M.1.1.3b,c). This may result in the forma-tion of a mud- or water-filled “blister”(Fig.M.1.1.3d,e,f). Shell coverage can also oc-cur over irritants attached to, or lying up againstthe inner shell, a process that may result in a“blister pearl” (Fig.M.1.1.3g).

Where perforation or other irritants exceed re-pair, the health of the mollusc is jeopardisedand it becomes susceptible to opportunistic in-fections (Fig.M.1.1.3.h). The degree of shellperforation can be determined by holding theshell up to a strong light.

Where abnormalities occurring within the ma-trix of the shell warrant further investigation,freshly collected specimens can be sent intactto the laboratory or fixed for subsequent de-calcification, as required.

M.1.1.4 Soft-Tissue Surfaces (Level I)

The appearance of the soft-tissues is frequentlyindicative of the physiological condition of theanimal. Gross features which should be re-corded include:• condition of the animal as listed below:

fat - the soft-tissues fill the shell, are turgidand opaquemedium - the tissues are more flaccid,opaque and may not fill the shell cavitywatery - the soft-tissues are watery/trans-parent and may not fill the shell cavity (Fig.(Fig.M.1.1.4a, Fig.M.1.1.4b)

• colour of the digestive gland – e.g., pale,mottled, dark olive

• any abnormal enlargement of the heart orpericardial cavity – e.g., cardiac vibriosis,tumours

• presence of focal lesions such as:

abnormal coloration (eg., patches of green,pink, red, black, etc.)abscesses (Fig. M.1.1.4c)tumour-like lesions (Fig. M.1.1.4d)tissue (e.g. gill) erosion

• presence of water blisters in the viscera, palp,or mantle (Fig.M.1.1.4e)

• presence of pearls or other calcareous de-posits (Fig.M.1.1.4f) within the soft tissues

• presence of parasites or commensals suchas:

pea crabs in mantle cavityparasitic copepods attached to gillspolychaetes, nematodes and turbellarians inmantle cavity or on surrounding surfaces(Fig. M.1.1.4g)redworm (Mytilicola spp.) usually exposedonly on dissection of the digestive tractciliates (sessile or free-swimming) and otherprotistans (for larvae only)bacteria (for larvae only)

• any mechanical (e.g., knife) damage to thesoft-tissues during the opening of the shell.

Abscess lesions, pustules, tissuediscolouration, pearls, oedema (water blisters),overall transparency or wateriness, gill defor-mities, etc., can be present in healthy molluscs,but, if associated with weak or dying animals,should be cause for concern. Record the lev-els of tissue damage and collect samples ofboth affected and unaffected animals for labo-ratory examination. Moribund animals, or thosewith foul-smelling tissues may be of little usefor subsequent examination (especially fromwarm water conditions), however, numbers af-fected should be recorded.

Worms or other organisms (e.g. pea crabs,copepods, turbellarians) on the soft-tissues arealso common and not generally associated withdisease. If present in high numbers on weakmolluscs, however, numbers should be notedand samples of intact specimens collected forlaboratory examination and identification. Fixa-tion in 10% buffered formalin is usually ad-equate for preserving the features necessaryfor subsequent identification.

M.1.2 Environmental Parameters(Level I)

Environmental conditions have a significant ef-fect on molluscan health, both directly (withinthe ranges of physiological tolerances) and in-directly (enhancing susceptibility to infections).This is especially important for species grownunder conditions which differ significantly fromthe wild (e.g. oysters grown in suspension).Important environmental factors for molluscanhealth include water temperature, salinity, tur-bidity, fouling and plankton blooms. Extremesand/or rapid fluctuations in these can seriouslycompromise molluscan health. Anthropogenicfactors include a wide range of biologic andchemical pollutants. Since molluscs are, essen-tially, sessile species (especially under cultureconditions) this renders them particularly sus-ceptible to pollution. In addition, molluscs have

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(SE McGladdery)

Fig.M.1.1.4e. Water blister (oedema/edema) inthe soft-tissues of the mantle margin of anAmerican oyster (Crassostrea virginica).

(SE McGladdery)

Fig.M.1.1.4f. Calcareous deposits (“pearls”) inthe mantle tissues of mussels in response toirritants such as mud or digenean flatwormcysts.

(SE McGladdery and M Stephenson)

Fig.M.1.1.4g. Polydoriid tunnels underlying thenacre at the inner edge of an American oyster(Crassostrea virginica) shell, plus another free-living polychaete, Nereis diversicolor on the in-ner shell surface.

(SE McGladdery)

Fig.M.1.1.4b. Watery oyster (Crassostreavirginica) tissues – compare with M.1.1.4a.

(SE McGladdery)

Fig.M.1.1.4c. Abscess lesions (creamy-yellowspots, see arrows) in the mantle tissue of a Pa-cific oyster (Crassostrea gigas).

(MS Park and DL Choi)

(SE McGladdery)

Fig.M.1.1.4a. Normal oyster (Crassostreavirginica) tissues.

Fig.M.1.1.4d. Gross surface lesions in Pacificoyster (Crassostrea gigas) due to Marteiliodeschungmuensis.

>

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low tolerance of some other water-uses/abuses(e.g., dynamite and cyanide fishing; dragging;creosote and other anti-fouling chemical com-pounds; agricultural run-off).

Maintaining records of temperature, salinity (inestuarine or coastal areas), turbidity and man-made disturbances provide valuable backgrounddata, essential for accurate interpretation of mor-tality observations and results from laboratoryanalyses.

M.1.3 General Procedures

M.1.3.1 Pre-Collection Preparation

Wherever possible, check the number of speci-mens required for laboratory examination withlaboratory personnel before collecting thesample. Ensure that each specimen is intact,i.e., no empty or mud-filled shells. Largersample numbers are generally needed forscreening purposes compared with numbersrequired for disease diagnosis.

M.1.3.2 Background Information (Level I)

All samples being submitted for laboratory ex-amination should include as much backgroundinformation as possible, such as:

• reason(s) for submitting the sample (mortalities, abnormal growth/spawning, healthscreening, etc.);

• gross observations and environmental paraeters (as described under M.1.1 and M.1.2);

• where samples are submitted due to mortalties, approximate prevalences and patterns ofmortality (acute or chronic/sporadic cumula-tive losses), and

• whether or not the molluscs are from local popu-lations or from another site. If the stock is notlocal, the source and date of transfer shouldalso be noted.

The above information will help identify if han-dling stress, change of environment or infec-tious agents may be a factor in mortalities. Itwill also help speed up accurate diagnosis of adisease problem or disease-risk analysis.

M.1.3.3 Sample Collection for Health Sur-veillance

The most important factors associated with col-lection of specimens for surveillance are:• sample numbers that are high enough (see

Table M.1.3.3 below)

• susceptible species are sampled• samples include age- or size-groups that are

most likely to manifest detetctable infections.Such information is given under specific dis-ease sections.

The standard sample sizes for screening healthyaquatic animals, including molluscs, is given inTable M.1.3.3 below.

M.1.3.4 Sample Collection for Disease Di-agnosis

All samples submitted for disease diagnosisshould include as much supporting informationas possible including:

• reason(s) for submitting the sample (mortali-ties, abnormal growth, etc.)

• handling activities (de-fouling, size sorting/grading, site changes, new species/stock in-troduction, etc.)

• history and origin(s) of the affectedpopulation(s);

• environmental changes

M.1.3.5 Live Specimen Collection for Ship-ping (Level I)

Once the required number of specimens hasbeen determined, and the laboratory has provideda date or time for receipt of the sample, the mol-luscs should be collected from the water. Thisshould take place as close to shipping as pos-sible to reduce air-storage changes in tissues andpossible mortalities during transportation. This isespecially important for moribund or diseasedmollusc samples.

The laboratory should be informed of the esti-mated time of arrival to ensure they have thematerials required to process the sample pre-pared before the sample arrives. This helps re-duce the time between removal from the waterand preservation of the specimens for exami-nation.

The molluscs should be wrapped in papersoaked with ambient seawater. For small seed(<10 mm), these can be packed in paper orstyrofoam cups along with damp paper towelto prevent movement during transportation.Larger molluscs should be shipped in insulatedand sealable (leakproof) coolers (styrofoam orplastic). Where more than one sample is in-cluded in the same cooler, each should beplaced in a separate and clearly labeled plasticbag (tied or ziploc). Use of plastic bags is re-quired to prevent exposure of marine species


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to freshwater ice (gel-paks or plastic bottlescontaining frozen water are recommended overloose ice to keep specimens cool) and to re-duce loss of mantle fluids.

Label containers clearly:

“Live Specimens, Store at _____°C to ______°CDO NOT FREEZE”

If being shipped by air also indicate:

“HOLD AT AIRPORT AND CALL FOR PICK-UP”

Clearly indicate the name and telephone num-ber of the contact person responsible for pick-ing up the package at the airport or receiving itat the laboratory.Ship early in the week to avoid arrival duringthe weekend with possible loss of samples dueto improper storage. Inform the contact personas soon as the shipment has been sent and,where appropriate, give them the name of thecarrier and waybill number.

1 Ossiander, F.J. and G. Wedermeyer. 1973. Journal of Fisheries Research Board of Canada 30:1383-1384.

M.1.3.6 Preservation (Fixation) of TissueSamples (Level I - with basic training)

For samples that cannot be delivered live to adiagnostic laboratory, due to distance or slowtransportation, specimens should be fixed (pre-served) on site. This is suitable for subsequenthistology examination, but means that routinebacteriology, mycology or media culture (e.g.,Fluid Thioglycollate Medium culture of Perkinsusspp.) cannot be performed. Diagnostic needsshould, therefore, be discussed with laboratorypersonnel prior to collecting the sample.

The following fixatives can be used for preser-vation of samples:

i) 1G4F solution (1% Glutaraldehyde : 4% Form-aldehyde)

*Stock 1G4F solution - may be held at 4oC forup to 3 months:

120 ml 37-40% buffered formalin solution**

20 ml 50% glutaraldehyde360 ml tap water

**Buffered formalin solution:1 litre 37-40% formaldehyde15 gm disodium phosphate (Na

2HPO

4)


Table M.1.3.31 . Sample sizes needed to detect at least one infected host in a population of agiven size, at a given prevalence of infection. Assumptions of 2% and 5% prevalences are mostcommonly used for surveillance of presumed exotic pathogens, with a 95% confidence limit.

Prevalence (%)

Population Size 0.5 1.0 2.0 3.0 4.0 5.0 10.0

50 46 46 46 37 37 29 20

100 93 93 76 61 50 43 23

250 192 156 110 75 62 49 25

500 314 223 127 88 67 54 26

1000 448 256 136 92 69 55 27

2500 512 279 142 95 71 56 27

5000 562 288 145 96 71 57 27

10000 579 292 146 96 72 29 27

100000 594 296 147 97 72 57 27

1000000 596 297 147 97 72 57 27

>1000000 600 300 150 100 75 60 30

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0.06 gm sodium hydroxide (NaOH)0.03 gm phenol red (pH indicator)

Working solution – should be preparedimmediately prior to use:

500 ml filtered ambient seawater orInstant Ocean

500 ml Stock 1G4F solution*

The required tissue thickness is about 2-3 mm.Tissues can tolerate long storage in this fixativeat room temperature. (N.B. Thicker tissues, orwhole animals, may be fixed using the 10%buffered formalin solution as described below).

ii) 10% Buffered formalin in filtered ambientseawater (This is the easiest solution to prepareand store).

10 ml 37-40% buffered formalinsolution**

90 ml filtered ambient seawater

N.B. Whole specimens less than 10 mm thickcan be fixed with this solution. If the specimensare larger, cut them into two or more piecesbefore fixing (ensure that pieces from differentspecimens do not get mixed up).

iii) Davidson’s Fixative

Tissue up to 10 mm in thickness can be fixedin Davidson’s fixative. Prior to embedding, tis-sues need to be transferred to either 50% etha-nol for 2 hours (minimum) and then to 70%ethanol, or directly to 70% isopropanol. Bestresults are obtained if fixative is made up in thefollowing order of ingredients.

Stock Solution:

400 ml glycerin800 ml formalin(37-40% formaldhyde)1200 ml 95% ethanol (or 99% iso propanol)1200 ml filtered natural or artificial

seawater

Working Solution: dilute 9 parts stock solutionwith 1 part glacial acetic acid

Important Notes:

• All fixatives should be kept away from openwater and used with caution against contactwith skin and eyes.

• If the molluscs cannot be fixed intact, con-tact the diagnostic laboratory to get guidancefor cracking shell hinges or removing the re-quired tissues.

M.1.3.7 Shipping Preserved Samples(Level I)

Many transport companies (especially air car-riers) have strict regulations regarding shippingany chemicals, including fixed samples for di-agnostic examination. Check with the carrierbefore collecting the sample to prevent loss oftime and/or specimens due to inappropriatepackaging, labeling, etc.. If the tissues havebeen adequately fixed (as described in M.1.3.4),most fixative or storage solution can be drainedfrom the sample for shipping purposes. As longas sufficient solution is left to keep the tissuesfrom drying out, this will minimise the quantityof chemical solution being shipped. Pack fixedsamples in a durable, leak-proof container.

Label containers clearly with the information asdescribed for live specimens (M.1.3.3.). Clearlyindicate the name and telephone number of thecontact person responsible for picking up thepackage at the airport or receiving it at the labo-ratory. Ship early in the week to avoid arrivalduring the weekend with possible loss ofsamples due to improper storage. Inform thecontact person as soon as the shipment hasbeen sent and, where appropriate, give themthe name of the carrier and waybill number.

If being shipped by air also indicate:

“HOLD AT AIRPORT AND CALL FOR PICK-UP”

M.1.4 Record-Keeping (Level I)

Record keeping is essential for effective dis-ease management. For molluscs, many of thefactors that should be recorded are outlined insections M.1.4.1, M.1.4.2, and M.1.4.3.

M.1.4.1 Gross Observations (Level I)

Gross observations can be included with rou-tine monitoring of mollusc growth, either bysub-sampling from suspension cages, lines orstakes, or by guess estimates from surfaceobservations.

For hatchery operations, the minimum essen-tial information which should be recorded/logged are:• feeding activity• growth• mortalities

These observations should be recorded on adaily basis for larval and juvenile molluscs, in-cluding date, time, tank, broodstock (where


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there are more than one) and food-source (al-gal culture batch or other food-source). Datesand times for tank and water changes shouldalso be noted, as well as dates and times forpipe flushing and/or disinfection. Ideally, theselogs should be checked regularly by the per-son responsible for the site/animals.

For open-water mollusc sites, the minimum es-sential observations which need to be recorded/logged include:

• growth• fouling• mortalities

These should be recorded with date, site loca-tion and any action if taken (e.g., defouling orsample collection for laboratory examination).Ideally, these logs should be checked regularlyby the person responsible for the site/animals.

M.1.4.2 Environmental Observations(Level I)

This is most applicable to open water sites, butshould also be included in land-based systemswith flow-through or well-based water sources.The minimum essential data which should berecorded are:

• temperature• salinity• turbidity (qualitative evaluation or secchi disc)• algal blooms• human activity

The frequency of these observations will varywith site. Where salinity or turbidity rarely vary,records may only be required during rainy sea-sons or exceptional weather conditions. Tem-perate climates will require more frequent wa-ter temperature monitoring than tropical cli-mates. Human activity should be logged on an“as it happens” basis for reference if no infec-tions or natural environmental changes can beattributed to a disease situation.

M.1.4.3 Stocking Records (Level I)

Information on movements of molluscs into andout of a hatchery should be recorded. Thisshould include:• exact source of the broodstock/seed• condition on arrival• date, time and person responsible for receiv-

ing delivery of the stock• date, time and destination of stock shipped

out of the hatchery

Where possible, animals from different sourcesshould not be mixed.

All movements of molluscs onto and off anopen-water site should also be recorded, in-cluding:

• exact source of the molluscs• condition on arrival• date, time and person responsible for receiv-

ing delivery of the stock• date, time and destination of stock shipped

off site

In addition, all movements of stocks within ahatchery, nursery or grow out site should belogged with the date for tracking purposes if adisease situation arises.

M.1.5 References

Elston, R.A. 1989. Bacteriological methods fordiseased shellfish, pp. 187-215. In: Austin,B. and Austin, D.A. (eds.) Methods for theMicrobiological Examination of Fish andShellfish. Ellis Horwood Ser. Aquac. Fish.Sup.. Wiley and Sons, Chichester, UK.

Elston, R.A. 1990. Mollusc diseases: Guide forthe Shellfish Farmer. Washington Sea GrantProgram, University of Washington Press,Seattle. 73 pp.

Elston, R.A., E.L. Elliot, and R.R. Colwell. 1982.Conchiolin infection and surface coatingVibrio: Shell fragility, growth depression andmortalities in cultured oysters and clams,Crassostrea virginica, Ostrea edulis andMercenaria mercenaria. J. Fish Dis. 5:265-284.

Fisher, W.S. (ed.). 1988. Disease Processes inMarine Bivalve Molluscs. Amer. Fish. Soc.Spec. Public. 18. American Fisheries Soci-ety, Bethesda, Maryland, USA.

Howard, D.W. and C.S. Smith. 1983. Histologi-cal Techniques for Bivalve Mollusks. NOAATech. Memo. NMFS-F/NEC-25.Woods Hole,Massachusetts. 97 pp.

Lauckner, G. 1983. Diseases of Mollusca:Bivalvia, pp. 477-520. In: O. Kinne(ed.) Dis-eases of Marine Animals Volume II. Introduc-tion Bivalvia to Scaphopoda. BiologischeAnstalt Helgoland, Hamburg.

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Luna, L.G. 1968. Manual of Histologic StainingMethods of the Armed Forces Institute of Pa-thology. McGraw-Hill Book Company, NewYork. 258 pp.

McGladdery, S.E., R.E. Drinnan, and M.F.Stephenson. 1993. A manual of the parasites,pests and diseases of Canadian Atlanticbivalves. Can. Tech. Rep. Fish. Aquat.Sci.1931.121 pp.

Ossiander, F.J. and G. Wedermeyer. 1973.Computer program for sample size requiredto determine disease incidence in fish popula-tions. J. Fish. Res. Bd. Can. 30: 1383-1384.

Pass, D.A., R. Dybdahl, and M.M. Mannion.1987. Investigations into the causes of mor-tality of the pearl oyster (Pinctada maxima)(Jamson), in Western Australia. Aquac.65:149-169.

Perkins, F.O. 1993. Infectious diseases ofmoluscs, pp. 255-287. In: Couch, J.A. andFournie, J.W. (eds.). Pathobiology of Marineand Estuarine Organisms. CRC Press, BocaRaton, Florida.


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M.2.1 Background Information

M.2.1.1 Causative Agents

Bonamiosis (a.k.a. Microcell Disease;haemocyte disease of flat or dredge oysters) iscaused by two Protistan (= Protozoan = single-celled) species belonging to the Haplosporidia:Bonamia ostreae and Bonamia sp.. More infor-mation about the disease can be found in theOIE Diagnostic Manual for Aquatic Animal Dis-eases (OIE 2000a).

M.2.1.2 Host Range

Bonamia ostreae occurs naturally in Ostrea edulis(European oyster) and O. conchaphila (O. lurida)(Olympia oyster). Other ostreiid species can be-come infected if transferred to enzootic areas,namely O. puelchana, O. angasi and Ostrealutaria (Tiostrea lutaria) (New Zealand oyster),Tiostrea chilensis (Ostrea chilensis) (SouthAmerican oyster), thus, all species of Ostrea,Tiostrea and some Crassostrea (C. ariakensis)should be considered susceptible. To date,Crassostrea gigas (Pacific oyster), Mytilus edulisand M. galloprovincialis (edible mussels) andRuditapes decussatus and R. philippinarum (Eu-ropean and Manila clams) have been found tobe resistant to infection. These species have alsobeen shown to be incapable of acting as reser-voirs or sub-clinical carriers of infection.

M.2.1.3 Geographic Distribution

Bonamia ostreae: The Netherlands, France,Spain, Italy, Ireland, the United Kingdom (exclud-ing Scotland) and the United States of America(States of California, Maine and Washington).Denmark, although stocked with infected oystersin the early 1980’s, has shown no sign of persis-tence of the infection and their European oystersare now considered to be free of B. ostreae.

Bonamia sp.: Australia (Western Australia,Victoria and Tasmania) and New Zealand (SouthIsland and southern North Island).

M.2.1.3 Asia-Pacific Quarterly Aquatic Ani-mal Disease Reporting System (1999 to2000)

For the reporting year 1999, Bonamia sp. waspositively reported in Australia in April, July andOctober in Tasmania; July and October in West-ern Australia. For the year 2000, Bonamia sp.was reported in March and April in Western Aus-tralia. In New Zealand, Bonamia sp. was

MOLLUSCAN DISEASESM.2 BONAMIOSIS

(BONAMIA SP., B. OSTREAE)

reported every month for 1999 and 2000reporting periods (OIE 1999, OIE 2000b).

M.2.2 Clinical Aspects

Most infections show no clinical signs until theparasites have proliferated to a level that elicitsmassive blood cell (haemocyte) infiltration anddiapedesis (Fig.M.2.2a). The pathology of in-fection varies with the species of Bonamia, andwith host species. Bonamia ostreae infects thehaemocytes of European oysters (Fig.M.2.2b),where it divides until the haemocyte bursts, re-leasing the parasites into the haemolymph. In-fections likely occur through the digestive tract,but gill infections suggest this may also be an-other infection route and macroscopic gill lesionsare sometimes visible. The pathology of Bonamiasp. in Australian Ostrea angasi and New Zealandpopulations of Tiostrea chilensis is very differ-ent. In Australia’s O. angasi, the first indicationof infection is high mortality. Surviving oystersrapidly start to gape on removal from the waterand may have “watery” tissues and a raggedappearance to the margin of the gill(unpublished data, B. Jones, Fisheries WesternAustralia). Bonamia sp. infects the walls of thegills, digestive ducts and tubules (Fig.M.2.2c),from which the parasites may be released intothe gut or surrounding water. Infectedhaemocytes may contain up to 6 Bonamia para-sites (Fig.M.2.2d). Infections induce massiveabscess-like lesions (haemocytosis), even in thepresence of only a few parasites. In T. chilensis,Bonamia sp. appears to enter via the gut wall(Fig.M.2.2e), and then infects the haemocytes,where up to 18 Bonamia per haemocyte can befound (Fig.M.2.2f). The resultant haemocytosisis less severe than in O. angasi. When infectedhaemocytes enter the gonad of T. chilensis toreabsorb unspawned gametes, the parasites pro-liferate and may be released via the gonoduct.Alternative release is also possible via tissuenecrosis following the death of the host. Despitethe differences in pathology, gene sequencingstudies (unpublished data, R. Adlard, Universityof Queensland, Australia) have shown that theAustralian and New Zealand Bonamia sp. are thesame species.

M.2.3 Screening Methods

More detailed methods for screening can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.

122

(SE McGladdery)

Fig.M.2.2a. Haemocyte infiltration and diaped-esis across intestinal wall of a European oyster(Ostrea edulis) infected by Bonamia ostreae.

(SE McGladdery)

Fig.M.2.2b. Oil immersion of Bonamia ostreaeinside European oyster (Ostrea edulis)haemocytes (arrows). Scale bar 20 µm.

(PM Hine)

Fig.M.2.2c. Systemic blood cell infiltration inAustralian flat oyster (Ostrea angasi) infectedby Bonamia sp. Note vacuolised appearanceof base of intestinal loop and duct walls (H&E).

M.2 Bonamiosis(Bonamia sp., B. ostreae)

(PM Hine)

Fig.M.2.2d. Oil immersion of Bonamia sp. in-fecting blood cells and lying free (arrows) in thehaemolymph of an infected Australian flat oys-ter, Ostrea angasi. Scale bar 20µm (H&E).

(PM Hine)

Fig.M.2.2e. Focal infiltration of haemocytesaround gut wall (star) of Tiostrea lutaria (NewZealand flat oyster) typical of infection byBonamia sp. (H&E).

(PM Hine)

Fig.M.2.2f. Oil immersion of haemocytespacked with Bonamia sp. (arrows) in an infectedTiostrea lutaria (H&E). >

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M.2.3.1 Presumptive

M.2.3.1.1 Gross Observations (Level I).

Slowed growth, presence of gill lesions (in somecases), gaping and mortalities of Ostrea edulisshould be considered suspect for Bonamiosis.Gross signs are not disease specific and requireLevel II examination.

M.2.3.1.2 Cytological Examination(Level II)

Spat or heart (preferably ventricle) smears or im-pressions (dabs) can be made onto a clean mi-croscope slide and air-dried. Once dry, the prepa-ration is fixed in 70% methanol. Quick and effec-tive staining can be achieved using commerciallyavailable blood-staining (cytological) kits, follow-ing the manufacturer’s instructions. The stainedslides are then rinsed (gently) in tapwater, allowedto dry and cover-slipped using a synthetic resinmounting medium. The parasite has basophilic(or colourless - Bonamia sp. in O. angasi) cyto-plasm and an eosinophilic nucleus (dependingon the stain used). An oil immersion observationtime of 10 mins per oyster preparation is consid-ered sufficient for screening cytology, tissue im-print and histology preparations (OIE 2000a).

M.2.3.2 Confirmatory

M.2.3.2.1 Histopathology (Level II)

It is recommended that at least two dorso-ven-tral sections through the cardiac cavity, gonadand gills of oysters over 18 months – 2 years (>30 mm shell height) be examined for screeningpurposes. These sections should be fixed imme-diately in a fast fixative such as 1G4F. Fixativessuch as Davidsons or 10% seawater bufferedformalin may be used for whole oysters (seeM.1.3.3.3), but these do not allow serial tissuesections to be collected for subsequent confir-matory Electron Microscopy (EM) diagnosis, ifrequired. Davidson’s fixative is recommendedfor subsequent PCR-based confirmation tech-niques.

Several standard stains (e.g., haematoxylin-eosin) enable detection of Bonamia spp.. Theparasites measure 2-5 µm and occur within thehaemocytes or epithelia (as described above) or,more rarely, loose within the haemolymph or gut/mantle lumens.

M.2.4 Diagnostic Methods

More detailed methods for diagnosis can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.

M.2.4.1 Presumptive

M.2.4.1.1 Histopathology and Cytology(Level II)

Histology and cytology (Level II), as describedunder M.2.3.2.1, may be used. For first-timediagnoses, a back up tissue specimen fixed forEM is recommended (M.2.4.2.1).

M.2.4. 2 Confirmatory

M.2.4.2.1 Transmission Electron Micros-copy (TEM) (Level III)

Tissue for TEM can be fixed in 1G4F (M.1.3.3.3),however, where it is likely that TEM will be re-quired for confirmatory diagnosis (M.2.4.1.1),small (< 1 mm cubed) sub-samples of infectedtissue should be fixed in 2-3% buffered glutaral-dehyde prepared with ambient salinity filteredseawater. Fixation should not exceed 1 hr.Longer storage in gluteraldehyde fixative is pos-sible, however membrane artifacts can be pro-duced. Tissues should be rinsed in a suitablebuffer prior to post-fixing in 1-2% osmium tetrox-ide (= osmic acid - highly toxic). This post-fixa-tive must also be rinsed with buffered filtered(0.22 µm) seawater prior to dehydration andresin-embedding.

Post-fixed tissues should be stored in a compat-ible buffer or embedded post-rinsing in a resinsuitable for ultramicrotome sectioning. Screen-ing of 1 micron sections melted onto glass mi-croscope slides and stained with Toluidine Blueis one method of selecting the tissue specimensfor optimum evidence of putative Bonamia spp.Ultrathin sections are then mounted on coppergrids (with or without formvar coating) for stain-ing with lead citrate + uranyl acetate or equiva-lent EM stain.

Ultrastructural differences between B. ostreaeand Bonamia sp. include diameter (B. ostreae =2.4 ± 0.5 µm; Bonamia sp. = 2.8 ± 0.4 µm in O.angasi, 3.0 ± 0.3 µm in T. chilensis); mean num-ber of mitochondrial profiles/section (B. ostreae= 2 ± 1; Bonamia sp. = 4 ± 1 in O. angasi, 3 ±1in T. chilensis), mean number ofhaplosporosomes/section (B. ostreae = 7 ± 5;

124

Bonamia sp. = 10 ± 4 in O. angasi, 14 ± 6 in T.chilensis); percentage of sections containinglarge lipid globules (B. ostreae = 7%; Bonamiasp. in O. angasi = 30%; in T. chilensis = 49%),large lipid globules/section (B. ostreae = 0.3 ±0.6; Bonamia sp. = 0.5 ± 0.8 in O. angasi, 0.8 ±0.9 in T. chilensis). Both species are distin-guished from Mikrocytos spp. by having a cen-trally-placed nucleus.

Plasmodial forms of Bonamia sp. in T. chilensisare distinguished from Bonamia ostreae by theirsize (4.0 -4.5 µm diameter), irregular cell andnucleus profile, amorphous cytoplasmic inclu-sions (multi-vesicular bodies) and arrays of Golgi-like smooth endoplasmic reticula. Other devel-opmental stages are more electron dense andsmaller in diameter (3.0 -3.5 µm).

M.2.5 Modes of Transmission

Prevalence and intensity of infection tends to in-crease during the warm water season withpeaks in mortality in September/October in thenorthern hemisphere, and January to April, inthe southern hemisphere. The parasite is diffi-cult to detect prior to the proliferation stage ofdevelopment or in survivors of an epizootic. Co-habitation and tissue homogenate/haemolymphinoculations can precipitate infections indicat-ing that transmission is direct (no intermediatehosts are required). There is a pre-patent pe-riod of 3-5 months between exposure and ap-pearance of clinical signs of B. ostreae infec-tion. In New Zealand the pre-patent period forBonamia sp. infection may be as little as 2.5months and rarely exceeds 4 months.

M.2.6 Control Measures

None known. Reduced stocking densities andlower water temperatures appear to suppressclinical manifestation of the disease, however,no successful eradication procedures haveworked to date. Prevention of introduction ortransfer of oysters from Bonamia spp. enzooticwaters into historically uninfected waters is rec-ommended.

M.2.7 Selected References

Adlard, R.D. and R.J.G. Lester. 1995. Develop-ment of a diagnostic test for Mikrocytosroughleyi, the aetiological agent of Australianwinter mortality in the commercial rock oys-ter, Saccostrea commercialis (Iredale &Roughley). J. Fish Dis. 18: 609-614.

Balouet, G., M. Poder, and A. Cahour. 1983.


Haemocytic parasitosis: Morphology and pa-thology of lesions in the French flat oyster,Ostrea edulis L. Aquac. 43: 1-14.

Banning, P. van 1982. The life cycle of the oys-ter pathogen Bonamia ostreae with a pre-sumptive phase in the ovarian tissue of theEuropean flat oyster, Ostrea edulis. Aquac.84: 189-192.

Carnegie, R.B., B.J. Barber, S.C. Culloty, A.J.Figueras, and D.L. Distel. 2000. Developmentof a PCR assay for detection of the oysterpathogen Bonamia ostreae and support forits inclusion in the Haplosporidia. Dis. Aquat.Org. 42(3): 199-206.

Culloty, S.C., B. Novoa, M. Pernas, M.Longshaw, M. Mulcahy, S.W. Feist, and A.J.

Figueras.1999. Susceptibility of a number ofbivalve species to the protozoan parasiteBonamia ostreae and their ability to act as vec-tors for this parasite. Dis. Aquat. Org. 37(1):73-80.

Dinamani, P., P.M. Hine, and J.B. Jones. 1987.Occurrence and characteristics of thehemocyte parasite Bonamia sp. on the NewZealand dredge oyster Tiostrea lutaria. Dis.Aquat. Org. 3: 37-44.

Farley, C.A., P.H. Wolf, and R.A. Elston. 1988.A long term study of “microcell” disease in oys-ters with a description of a new genus,Mikrocytos (g.n.) and two new species,Mikrocytos mackini (sp.n.) and Mikrocytosroughleyi (sp.n.). Fish. Bull. 86: 581-593.

Friedman, C.S. and F.O. Perkins. 1994. Rangeextensiion of Bonamia ostreae to Maine,U.S.A. J. Inverteb. Pathol. 64: 179-181.

Hine, P.M. 1991. Ultrastructural observations onthe annual infection pattern of Bonamia sp. inflat oysters, Tiostrea chilensis. Aquac. 93: 241-245.

McArdle, J.F., F. McKiernan, H. Foley, and D.H.Jones. 1991. The current status of Bonamiadisease in Ireland. Aquac. 93:273-278.

Mialhe, E., E. Bachere, D. Chagot, and H. Grizel.1988. Isolation and purification of the proto-zoan Bonamia ostreae (Pichot et al., 1980), aparasite affecting flat oyster Ostrea edulis L.Aquac. 71: 293-299.

OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35 p.

125

M.3 MARTEILIOSIS(MARTEILIA REFRINGENS,

M. SYDNEYI)



Marteiliosis is caused by two species of para-sites, belonging to the Phylum Paramyxea.Marteilia refringens is responsible for Aber Dis-ease (a.k.a Digestive Gland Disease) of Euro-pean oysters (Ostrea edulis) and Marteiliasydneyi is responsible for QX Disease ofSaccostrea glomerata (syn. Crassostreacommercialis, Saccostrea commercialis) and,possibly, Saccostrea echinata. More informa-tion about the disease can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000a).

M.3.1.2 Host Range

Ostrea edulis is infected by Marteilia refringens.Other host species include Tiostrea chilensis,Ostrea angasi, O. puelchana, Cerastoderma (=Cardium) edule, Mytilus edulis, M.galloprovincialis, Crassostrea gigas and C.virginica. Marteilia sydneyi infects Saccostreaglomerata and possibly S. echinata. Anothermarteiliad, Marteilia maurini, infects mussels(Mytilus edulis and M. galloprovincialis) fromFrance, Spain and Italy. This species is notreadily distinguished morphologically from M.refringens and distinct species status is underinvestigation. An unidentified marteiliad was re-sponsible for mass mortalities of the Calico scal-lop (Argopecten gibbus) in Florida in the late1980’s, but has not re-appeared since. AnotherMarteilia-like species was reported from the gi-ant clam, Tridacna maxima. Other species ofMarteilia that have been described include M.lengehi from Saccostrea (Crassostrea) cucullata(Persian Gulf and north Western Australia) andM. christenseni in Scrobicularia plana (France).These are differentiated from M. refringens andM. sydneyi by the cytoplasmic contents of thesporangia and spore morphology.


Marteilia refringens is found in O. edulis in south-ern England, France, Italy, Portugal, Spain, Mo-rocco and Greece. Marteilia sydneyi is found inS. glomerata in Australia (New South Wales,Queensland and Western Australia).

M.3.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System (1999-2000)

No positive report of the disease in any coun-try for the 2 year reporting periods. Most coun-tries have no information about the occurrenceof the disease (OIE 1999,OIE 2000b).


Early stages of Marteilia refringens develop in thedigestive ducts, intestinal and stomach epitheliaand gills (Fig.M.3.2a). Later, spore-formingstages appear in the blind-ending digestive tu-bule epithelia (Fig.M.3.2b). Proliferation of theparasite is associated with emaciation and ex-haustion of glycogen reserves, grossdiscolouration of the digestive gland, cessationof feeding and weakening. Mortalities appear tobe associated with sporulation of the parasite anddisruption of the digestive tubule epithelia.



M.3.3.1 Presumptive

M.3.3.1.1 Gross Observations (Level I)

Slowed growth, gaping and mortalities of Ostreaedulis and other susceptible species should beconsidered suspect for Marteiliosis. Gross signsare not specific for Bonamiosis or Marteiliosis andrequire Level II examination.

M.3.3.1.2 Tissue Imprints (Level II)

Cut a cross-section through the digestive gland,blot away excess water with blotting paper anddab the cut section of the digestive gland onto aclean microscope slide. Fix tissue imprint for 2-3min in 70% methanol. Quick and effective stain-ing can be achieved with a commercially avail-able blood-staining (cytological) kit, using themanufacturer’s instructions. The stained slidesare then rinsed (gently) under tap water, allowedto dry and cover-slipped using a synthetic resinmounting medium.

The parasite morphology is as described for his-tology (M.3.3.2.1), although colouration may varywith the stain chosen. Initial screening with ahaematoxylin or trichrome stain, as used for his-

126

M.3 Marteiliosis(Marteilia refringens, M. sydneyi)

(SE McGladdery)

Fig.M.3.2a. Digestive duct of a European oys-ter, Ostrea edulis, showing infection of distalportion of the epithelial cells by plasmodia (ar-rows) of Marteilia refringens. Scale bar 15 µm(H&E).

(SE McGladdery)

Fig.M.3.2b. Digestive tubule of a Europeanoyster, Ostrea edulis, showing refringent sporestage of Marteilia refringens (star). Scale bar 50µm (H&E).

(RD Adlard)

Fig.M.3.4.1.1a. Tissue imprint from Saccostreacommercialis (Sydney rock oyster) heavily in-fected by Marteilia sydneyi (arrows) (QX dis-ease). Scale bar 250 µm (H&E).

(RD Adlard)

Fig.M.3.4.1.1b. Oil immersion of tissue squashpreparation of spore stages of Marteilia sydneyifrom Sydney rock oyster (Saccostreacommercialis) with magnified inset showing twospores within the sporangium. Scale bar 50 µm(H&E).

tology, may assist familiarisation with tissue im-print characteristics prior to using a dip-quickmethod. An observation time of 10 mins at 10-25x magnification is considered sufficient forscreening purposes.



Two dorso-ventral tissue section (2-3 mm thick)are recommended for screening purposes. Thesecan be removed from oysters over 18-24 monthsold (or >30 mm shell height) for immediate fixa-tion in a fast fixative, such as 1G4F. Davidsonsor 10% buffered formalin may be used for largersamples or whole oysters (see M.1.3.3.3) butthese provide less satisfactory results if subse-quent processing for Transmission Electron Mi-croscopy (TEM) (M.3.4.2.1) is required (e.g., forspecies identification). Several standard stains(e.g., haematoxylin-eosin) enable detection ofMarteilia spp..

The early stages of development occur in thestomach, intestine and digestive duct epithelia(usually in the apical portion of the cell) and ap-pear as basophilic, granular, spherical inclu-sions (Fig.M.3.2a). Later stages occur in thedigestive tubules, where sporulation may in-duce hypertrophy of the infected cell. Marteiliaspp. spores contain eosinophilic, “refringent”,bodies which are easily detected at 10-25 xmagnification under light microscopy(Fig.M.3.2b).

127



M.3.4.1 Presumptive

M.3.4.1.1 Tissue Imprints (Level II)

As described under M.3.3.1.2, tissue imprintsmay also be used for presumptive diagnoses(Fig.M.3.4.1.1a,b). For first-time diagnoses backup tissues should be fixed for histology and EMconfirmatory diagnosis.


Histology techniques as described underM.3.3.2.1, may be used. For first-time diag-noses a back up tissue specimen fixed for EMis recommended, as described below.



TEM tissue preparation involves fixing tissueseither in 1G4F (M.1.3.3.3) or small (< 1 mmcubed) sub-samples of infected tissue in 2-3%glutaraldehyde mixed and buffered for ambientfiltered seawater. Ideally, fixation in 2-3%gluteraldehyde should not exceed 1 hr, sincelonger storage may induce membranous artifacts.Tissues should be fixed in 1G4F for 12-24 hrs.Following primary fixation, rinse tissues in a suit-able buffer and post-fix in 1-2% osmium tetrox-ide (OsO

4= osmic acid - highly toxic). Second-

ary fixation should be complete within 1 hr. TheOsO

4fixative must also be rinsed with buffer/fil-

tered (0.22 µm) seawater prior to dehydrationand resin-embedding.

Post-fixed tissues can be stored in a compatiblebuffer or embedded post-rinsing in a resin suit-able for ultramicrotome sectioning. Screening of1 µm sections melted onto glass microscopeslides with 1% toluidine blue solution is onemethod of selecting the tissue specimens foroptimum evidence of possible Marteilia spp. Ul-trathin sections are then mounted on copper grids(with or without formvar coating), and stained withlead citrate + uranyl acetate (or equivalent EMstain).


Marteilia refringens plasmodia contain striatedinclusions, eight sporangial primordia, with upto four spores to each mature sporangium.Marteilia sydneyi has a thick layer of concen-tric membranes surrounding the mature spore,lacks striated inclusions in the plasmodia, formseight to sixteen sporangial primordia in eachplasmodium and each sporangium containstwo (rarely three) spores.

M.3.4.2.2 In situ Hybridization (Level III)

In situ hybridization (Level III) techniques areunder development but not yet available com-mercially. Information on the current status ofthese and related molecular probe techniquesmay be obtained from IFREMER Laboratory atLa Tremblade, France (OIE 2000a, Annex MAI).


Marteilia refringens transmission appears to berestricted to periods when water temperaturesexceed 17°C. High salinities may impedeMarteilia spp. multiplication within the host tis-sues. Marteilia sydneyi also has a seasonal pe-riod of transmission with infections occurringgenerally from mid- to late-summer (January toMarch). Heavy mortalities and sporulation occurall year round. The route of infection and life-cycleoutside the mollusc host are unknown. Since ithas not been possible to transmit the infectionexperimentally in the laboratory, an intermediatehost is suspected. This is reinforced by recentobservations showing spores do not survive morethan 7-10 days once isolated from the oyster.Cold temperatures prolong survival (35 days at15°C). Spore survival within fish or birds was lim-ited to 2 hrs, suggesting they are an unlikely modeof dispersal or transmission.


None known. High salinities appear to suppressclinical manifestation of the disease, however,no eradication attempts have been successful,to date. Prevention of introduction or transfer ofoysters or mussels from Marteilia spp. enzooticwaters into historically uninfected waters is rec-ommended.


Anderson, T.J., R.D. Adlard, and R.J.G. Lester.1995. Molecular diagnosis of Marteilia sydneyi(Paramyxea) in Sydney rock oysters,Saccostrea commercialis (Angas). J. Fish Dis.18(6): 507-510.

128

Auffret, M. and M. Poder. 1983. Studies ofMarteilia maurini, parasite of Mytilus edulisfrom the north coasts of Brittany. Revue desTravaux de l’Institut des P?ches Maritimes,Nantes 47(1-2): 105-109.

Berthe, F.C.J., M. Pernas, M. Zerabib, P. Haffner,A. Thebault, and A.J. Figueras. 1998. Experi-mental transmission of Marteilia refringenswith special consideration of its life cycle. Dis.Aquat. Org. 34(2): 135-144.

Berthe, F.C.J., F. Le Roux, E. Peyretaillade, P.Peyret, D. Rodriguez, M. Gouy, and C.P.Vivares. 2000. Phylogenetic analysis of thesmall subunit ribosomal RNA of Marteiliarefringens validates the existence of PhylumParamyxea (Desportes and Perkins, 1990).J. Eukaryote Microbiol. 47(3): 288-293.

Comps, M. 1983. Morphological study of Marteiliachristenseni sp.n., parasite of Scrobiculariapiperata P. (Mollusc, Pelecypod). Revue desTravaux de l’Institut des P?ches MaritimesNantes 47(1-2): 99-104.

Hine, P.M. and T. Thorne. 2000. A survey of someparasites and diseases of several species ofbivalve mollusc in northern Western Austra-lia. Dis. Aquat. Org. 40(1): 67 -78.

Kleeman, S.N. and R.D. Adlard. 2000. Molecu-lar detection of Marteilia sydneyi, pathogen ofSydney rock oysters. Dis. Aquat. Org. 40(2):137-146.

Montes, J., M.A. Longa, A. Lama, and A. Guerra.1998. Marteiliosis of Japanese oyster(Crassostrea gigas) reared in Galicia NWSpain. Bull. Europ. Soc. Fish Pathol. 18(4):124-126.

Moyer, M.A., N.J. Blake, and W.S. Arnold. 1993.An ascetosporan disease causing mass mor-tality in the Atlantic calico scallop, Argopectengibbus (Linnaeus, 1758). J. Shellfish Res.12(2): 305-310.

Norton, J.H., F.O. Perkins, and E. Ledua. 1993.Marteilia-like infection in a giant clam, Tridacnamaxima, in Fiji. Journal of Invertebrate Pathol-ogy 61(3): 328-330.

OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.


OIE. 2000a. Diagnostic Manual for Aquatic Ani-mal Diseases, Third Edition, 2000. Office In-ternational des Epizooties, Paris, France.237p.

OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.

Renault, T., N. Cochennec, and B. Chollet. 1995.Marteiliosis in American oysters Crassostreavirginica reared in France. Dis. Aquat. Org.23(3): 161-164.

Robledo, J.A.F. and A. Figueras. 1995. The ef-fects of culture-site, depth, season and stocksource on the prevalence of Marteiliarefringens in cultured mussels (Mytilusgalloprovincialis) from Galicia, Spain. J.Parasitol. 81(3): 354-363.

Roubal, F.R., J. Masel, and R.J.G. Lester. 1989.Studies on Marteilia sydneyi, agent of QX dis-ease in the Sydney rock oyster, Saccostreacommercialis, with implications for its life-cycle.Aust. J. Mar. Fresh. Res. 40(2): 155-167.

Villalba, A., S.G. Mourelle, M.C. Lopez, M.J.Caraballal, and C. Azevedo. 1993. Marteiliasisaffecting cultured mussels Mytilusgalloprovincialis of Galicia (NW Spain). 1. Eti-ology, phases of the infection, and temporaland spatial variability in prevalence. Dis. Aquat.Org. 16(1): 61-72.

Villalba, A., S.G. Mourelle, M.J. Carballal, andC. Lopez. 1997. Symbionts and diseases offarmed mussels Mytilus galloprovincialisthroughout the culture process in the Rias ofGalicia (NW Spain). Dis. Aquat. Org. 31(2):127-139.

Wesche, S.J., R.D. Adlard, and R.J.G. Lester.1999. Survival of spores of the oyster patho-gen Marteilia sydneyi (Protozoa, Paramyxea)as assessed using fluorogenic dyes. Dis.Aquat. Org. 36(3): 221-226.

129



Mikrocytosis is caused by two species of para-sites of uncertain taxonomic affinity. Mikrocytosmackini is reponsible for Denman Island Disease(Microcell disease) of Pacific oysters(Crassostrea gigas), and Mikrocytos roughleyi isresponsible for Australian Winter Disease (Win-ter Disease, Microcell Disease) of Sydney rockoysters, Saccostrea glomerata. More informationabout the disease can be found in the OIE Diag-nostic Manual for Aquatic Animal Diseases (OIE2000a).

M.4.1.2 Host Range

Mikrocytos mackini naturally infects Crassostreagigas (Pacific oysters). Ostrea edulis (Europeanoysters), O. conchaphila (= O. lurida) (Olympiaoyster) and Crassostrea virginica (American oys-ters) growing in enzootic waters are also sus-ceptible to infection. Saccostrea glomerata(Crassostrea commercialis, Saccostreacommercialis) (Sydney rock oyster) is the onlyknown host for Mikrocytos roughleyi.


Mikrocytos mackini is restricted to specific locali-ties around Vancouver Island and southwestcoast of the Pacific coast of Canada. The para-site is limited to waters with temperatures below12?C. Mikrocytos roughleyi occurs in central tosouthern New South Wales, and at Albany andCarnarvon, Western Australia.


No positive report during the reporting periodsfor 1999 and 2000. In Australia, last year of oc-currence was 1996 (in New South Wales andWestern Australia. Most countries have no infor-mation about occurrence of the disease (OIE1999, OIE 2000b).


Mikrocytos mackini initiates focal infections of thevesicular connective tissue cells. This elicitshaemocyte infiltration and abscess formation.Grossly visible pustules (Fig.M.4.2a), abscesslesions and tissue ulcers, mainly in the mantle,may correspond to brown scar formation on the

M.4 MIKROCYTOSIS(MIKROCYTOS MACKINI,

M. ROUGHLEYI)

adjacent surface of the inner shell. However,such lesions are not always present. Small cells,1-3 µm in diameter, are found (rarely) aroundthe periphery of advanced lesions, or in con-nective tissue cells in earlier stages of diseasedevelopment. Severe infections appear to berestricted to oysters over 2 years old.

Mikrocytos roughleyi induces a systemic intrac-ellular infection of the haemocytes (never theconnective tissue cells) which may result in fo-cal lesions in the gills, connective, gonadal anddigestive tract.



M.4.3.1 Presumptive


Slowed growth, gaping and mortalities ofCrassostrea gigas and Saccostrea glomeratashould be considered suspect for Mikrocytosis.Gross signs are not pathogen specific and re-quire Level II examination, at least for first-timeobservations.

M.4.3.1.2 Cytological Examination andTissue Imprints (Level II)

Heart impressions (dabs) can be made onto aclean microscope slide and air-dried. Once dry,the slide is fixed in 70% methanol. Quick andeffective staining can be achieved with commer-cially available blood-staining (cytological) kits,using the manufacturer’s instructions. Thestained slides are then rinsed (gently) under tapwater, allowed to dry and cover-slipped using asynthetic resin mounting medium. Intracytoplas-mic parasites in the haemocytes will match thedescriptions given above for histology. This tech-nique is more applicable to M. roughleyi than M.mackini.

Tissue sections through mantle tissues (espe-cially abscess/ulcer lesions, where present) arecut and excess water removed with blotting pa-per. The cut section is dabbed onto a clean mi-croscope slide, fixed for 2-3 minutes in 70%methanol and stained. Quick and effective stain-ing can be achieved using a commercially avail-able blood-staining (cytological) kits, using themanufacturer’s instructions. The stained slides

130

M.4 MIKROCYTOSIS(Mikrocytos mackini, M. roughleyi)

(SM Bower)

Fig.M.4.2a. Gross abscess lesions (arrows) inthe mantle tissues of a Pacific oyster(Crassostrea virginica) severely infected byMikrocytos mackini (Denman Island Disease).

(SM Bower)

Fig.M.4.3.2.1a. Histological section throughmantle tissue abscess corresponding to thegross lesions pictured in Fig.M.4.2a, in a Pa-cific oyster (Crassostrea gigas) infected byMikrocytos mackini (H&E).

(SM Bower)

Fig.M.4.3.2.1b. Oil immersion of Mikrocytosmackini (arrows) in the connective tissue sur-rounding the abscess lesion pictured inFig.M.4.3.2.1a. Scale bar 20 µm (H&E).

are then rinsed (gently) under tap water, allowedto dry and cover-slipped using a synthetic resinmounting medium.

The parasite morphology is as described for his-tology (M.4.3.2.1), although colouration mayvary with the stain chosen. Initial screening witha haematoxylin or trichrome stain, as used forhistology, may assist familiarisation with tissueimprint characteristics prior to using a dip-quickmethod. An observation time of 10 mins underoil immersion is considered sufficient for screen-ing purposes.



It is recommended that at least two dorso-ven-tral sections (2-3 mm) through each oyster beexamined using oil immersion for screening pur-poses. Sections from oysters >2 yrs (or >30 mmshell height) should be fixed immediately in afast fixative such as 1G4F. Davidsons or 10%buffered formalin may be used for smaller orwhole oysters (see M.1.3.3.3) but these fixa-tives are not optimal for subsequent confirma-tory Electron Microscopy (EM) diagnosis(M.4.4.2.1), or species identification, if required.Also smaller oysters are not the recommendedsize-group for Mikrocytos screening. Sectionsthrough pustules, abscess or ulcer lesionsshould selected where present. Several stan-dard stains (e.g., haematoxylin-eosin) enabledetection of Mikrocytos spp.

Mikrocytos mackini appears as 2-3 µm intrac-ellular inclusions in the cytoplasm of the ve-sicular connective tissues immediately adjacentto the abscess-like lesions (Fig. M.4.3.2.1a,b).It may also be observed in muscle cells and,occasionally, in haemocytes or free, within thelesions. It is distinguished from Bonamia by aneccentric nucleus and from M. roughleyi by theconsistent absence of a cytoplasmic vacuole,and the presence of a mitochondrion in M.roughleyi. These features will not be clear un-der oil and require confirmation using 1 micronresin sections or TEM (described below). Nei-ther of these techniques, however, are practi-cal for screening purposes.

Mikrocytos roughleyi measures 1-3 µm in di-ameter and occurs exclusively in thehaemocytes. A cytoplasmic vacuole may ormay not be present. When present, it displacesthe nucleus peripherally. Nucleolar structuresmay or may not be visible under oil immersionresolution for this intracellular parasite.

131



M.4.4.1 Presumptive

M.4.4.1.1 Histopathology and Tissue Im-prints (Level II)

Histology (M.4.3.2.1) may be used, however, forfirst-time diagnoses, EM confirmation is recom-mended (M.4.4.2.2). Tissue imprints may also beused for presumptive diagnoses, where theydemonstrate the features described underM.4.3.1.2.


M.4.4.2.1 Histopathology and Tissue Im-prints (Level II)

Histology (M.4.3.2.1) and tissue imprints(M.4.3.1.2) may be used, however, for first-timediagnoses, EM confirmation is recommended(M.4.4.2.2).


Tissues should be fixed in 1G4F for 12-24 hours.Following primary fixation, rinse tissues in a suit-able buffer and post-fix in 1-2% osmium tetrox-ide (OsO4 = osmic acid - highly toxic). Second-ary fixation should be complete within 1 hour. TheOsO4fixative must also be rinsed with buffer/fil-tered (0.22 microns) seawater prior to dehydra-tion and resin-embedding.

Post-fixed tissues can be stored in a compatiblebuffer or embedded post-rinsing in a resin suit-able for ultramicrotome sectioning. Screening of1 micron sections melted onto glass microscopeslides with 1% toludine blue solution is onemethod of selecting the tissue specimens foroptimum evidence of putative Mikrocytos spp.Ultrathin sections are then mounted on coppergrids (with or without formvar coating), andstained with lead citrate + uranyl acetate (orequivalent EM stain).

Mikrocytos mackini is distinguished ultrastructur-ally (as well as by tissue location and host spe-cies) from Bonamia spp. by the location of thenucleolus. In M. mackini it is in the centre of thenucleus, while in B. ostreae it is eccentric.

Mikrocytos mackini also lacks mitochondria.The ultrastructural characteristics of Mikrocytosroughleyi have not been published, however, itis distinguished from M. mackini by the presenceof a cytoplasmic vacuole (along with completelydifferent geographic, host and tissue locations!).


Mikrocytos mackini transmission appears re-stricted to early spring (April-May) following pe-riods of 3-4 months at water temperatures <10˚C. High salinities (30-35 ppt) appear to favourparasite proliferation and mortalities of approxi-mately 40% occur in sub-tidal or low-tide popu-lations of older oysters.

Mikrocytos roughleyi is also associated with lowtemperatures and high salinities killing up to 70%of mature Sydney rock oysters in their third win-ter before marketing. This usually follows a pre-patent (sub-clinical) period of approximately 2.5months.

Transmission of M. mackini has been achievedby exposure of susceptible oysters tohomogenates from infected oysters as well as toproximal exposure, thus, it is believed that thisspecies has a direct life-cycle. M. roughleyi isalso thought to be transmitted directly from oys-ter to oyster.


Circumvention of mortalities has been achievedfor M. mackini at enzootic sites by relaying oys-ters to high tide levels during the peak transmis-sion period in April-May to reduce exposure tothe water-borne infectious stages. No controlmeasures are known for M. roughleyi.


Bower, S.M., D. Hervio, and S.E. McGladdery.1994. Potential for the Pacific oyster,Crassostrea gigas, to serve as a reservoir hostand carrier of oyster pathogens. ICES CouncilMeeting Papers, ICES, Copenhagen, Den-mark.1994. 5pp.

Bower, S.M. and G.R. Meyer. 1999. Effect ofcold-water on limiting or exacerbating someoyster diseases. J. Shellfish Res. 18(1): 296(abstract).

Farley, C.A., P.H. Wolf, and R.A. Elston. 1988.A long-term study of “microcell” disease in oys-ters with a description of a new genus,Mikrocytos (g. n.), and two new species,

M.4 Mikrocytosis(Mikrocytos mackini, M. roughleyi)

132

Mikrocytos mackini (sp. n.) and Mikrocytosroughleyi (sp. n.). Fish. Bull. 86(3): 581 -594.

Hervio, D., S.M. Bower, and G.R. Meyer. 1995.Life-cycle, distribution and lack of host speci-ficity of Mikrocytos mackini, the cause ofDenman Island disease of Pacific oysters(Crassostrea gigas). J. Shellfish Res. 14(1):228 (abstract).

Hervio, D., S.M. Bower, and G.R. Meyer. 1996.Detection, isolation and experimental trans-mission of Mikrocytos mackini, a microcellparasite of Pacific oysters Crassostrea gigas(Thunberg). J. Inverteb. Pathol. 67(1): 72-79.

Lester, R.J.G. 1990. Diseases of cultured mol-luscs in Australia. Advances in Tropical Aquac-ulture: Workshop, Tahiti, French Polynesia Feb.20 – Mar. 4, 1989. Actes de colloquesIFREMER 9: 207-216.




Smith, I.R., J.A. Nell, and R.D. Adlard. 2000. Theeffect of growing level and growing method onwinter mortality, Mikrocytos roughleyi, in dip-loid and triploid Sydney rock oysters,Saccostrea glomerulata. Aquac. 185(3-4): 197-205.

M.4 Mikrocytosis(Mikrocytos mackini, M. roughleyi)

133

M.5 PERKINSOSIS(PERKINSUS MARINUS, P. OLSENI)


P. marinus was not reported in Australia during1999 and 2000 reporting periods; P. olseni like-wise not reported in 1999 and 2000 (last year ofoccurrence in South Australia in 1997, and in1995 in New South Wales and Western Austra-lia). Suspected in Korea RO for reporting period1999 and 2000. In New Zealand, positively re-ported from April to December 2000. Perkinsusolseni occurs in wild populations of New Zealandcockles, Austrovenus stutchburyi (FamilyVeneridae) and two other bivalve species,Macomona liliana (Family Tellinidae) and Barbatianovae-zelandiae (Family Arcidae). These spe-cies occur widely in the coast of New Zealand.Affected locations have been the Waitemata andKaipara Harbours but the organism is probablyenzootic in the warmer waters of northern NewZealand (OIE 1999, OIE 2000a).


The effects of Perkinsus marinus on Crassostreavirginica range from pale appearance of the di-gestive gland, reduced condition indices, severeemaciation, gaping, mantle retraction, retardedgonadal development and growth and occasionalabscess lesions. Mortalities of up to 95% haveoccurred in infected C. virginica stocks.

Proliferation of Perkinsus olseni results in disrup-tion of connective and epithelial tissues and somehost species show occasional abscess forma-tion. Pustules up to 8 mm in diameter in affectedHaliotis spp. reduce market value and have beenassociated with heavy losses in H. laevi gata.



M.5.3.1 Presumptive


Slowed growth, gaping and mortalities ofCrassostrea virginica and Haliotis spp., as wellas other mollusc species in Perkinsus-enzooticwaters should be considered suspect forPerkinsiosis. Gross signs are not pathogen-spe-cific and require Level II examination, at least forfirst time observations.



Perkinsosis is caused by two species of protistanparasite belonging to the phylum Apicomplexa(although recent nucleic acid investigations sug-gest a possible affiliation with the dinoflagellates).Perkinsus marinus is responsible for “Dermo”disease in Crassostrea virginica (American oys-ters) and Perkinus olseni causes perkinsosis inmany bivalve species in tropical and subtropicalwaters. Other perkinsiid species are known toinfect clams in Europe (Perkinsus atlanticus) andthe eastern USA (Perkinsus spp.), as well asJapanese (Yesso) scallops, Patinopectenyessoensis in Pacific Canada (Perkinsusqugwadi). The taxonomic relationship betweenthese and the two species listed as ‘notifiable’ byOIE is currently under investigation. More infor-mation about the disease can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000a).

M.5.1.2 Host Range

Perkinsus marinus (formerly known asDermocystidium marinum and Labyrinthomyxamarinus) infects Crassostrea virginica (Americanoysters). Experimental infection to C. gigas (Pa-cific oysters) is possible, but they appear moreresistant than C. virginica. Perkinsus olsenishows a strong rDNA similarity to Perkinsusaltanticus of Ruditapes decussatus and the spe-ciation within this genus, as mentioned underM.5.1.1, is currently under nucleic acid investi-gation. Recognised hosts of P. olseni are theabalone species: Haliotis rubra, H. cyclobates,H. scalaris and H. laevigata. More than 50 othermolluscan species harbour Perkinsus spp., aswell as other possibly related species, withoutapparent harmful effects (e.g., in Arca clams[Fig.M.5.1.2a] and Pinctada pearl oysters[Fig.M.5.1.2b]).


Perkinsus marinus is found along the east coastof the United States from Massachusetts toFlorida, along the Gulf of Mexico coast to Ven-ezuela, and in Puerto Rico, Cuba and Brazil. Ithas also been introduced into Pearl Harbour,Hawaii. Range extension into Delaware Bay, NewJersey, Cape Cod and Maine are attributed torepeated oyster introductions and increased win-ter water temperatures. Perkinsus olseni occursin South Australia. Other species occur in Atlan-tic and Pacific oceans and the MediterraneanSea.

134



It is recommended that at least two dorso-ven-tral sections through each oyster be examinedusing oil immersion for screening purposes. Sec-tions from oysters >2 yrs (or >30 mm shell height)should be fixed immediately in a fast fixative suchas 1G4F. Davidson’s or 10% buffered formalinmay be used for smaller or whole oysters (seeM.1.3.3.3) but these fixatives are not optimal forsubsequent confirmatory Electron Microscopy(EM) diagnosis (M.5.4.2.1), or species identifi-cation, if required. Sections through pustules,abscess or ulcer lesions should be selected,where present. Several standard stains (e.g.,

M.5 Perkinsosis(Perkinsus marinus, P. olseni)

(SE McGladdery)

Fig.M.5.3.2.1b. Schizont (‘rosette’) stages ofPerkinsus marinus (arrows), the cause of‘Dermo’ disease in American oyster(Crassostrea virginica) digestive gland connec-tive tissue. Scale bar 30 µm (H&E).

(SE McGladdery)

Fig.5.3.2.2. Enlarged hypnospores of Perkinsusmarinus stained blue-black with Lugol’s iodinefollowing incubation in fluid thioglycollate me-dium. Scale bar 200 µm.

(PM Hine)

Fig.M.5.1.2a. Arca clam showing a Perkinsus-like parasite within the connective tissue. Mag-nified insert shows details of an advanced ‘sch-izont’ like stage with trophozoites showingvacuole-like inclusions. Scale bar 100 µm.(H&E).

(PM Hine)

Fig.M.5.1.2b. Pinctada albicans pearl oystershowing a Perkinsus-like parasite. Magnifiedinsert shows details of a ‘schizont’-like stagecontaining ‘trophozoites’ with vacuole-like in-clusions. Scale bar 250 µm (H&E).

(SM Bower)

Fig.M.5.3.2.1a. Trophozoite (‘signet-ring’)stages of Perkinsus marinus (arrows), the causeof ‘Dermo’ disease in American oyster(Crassostrea virginica) connective tissue. Scalebar 20 µm (H&E).

135

M.5.4.1.2 Fluid Thioglycollate MediumCulture (Level II)

Fluid thioglycollate medium culture (Level II) mayalso be used for presumptive diagnosis(M.5.3.2.2).



TEM is required to detect the species-specificultrastructure of the zoospore stage of develop-ment (collected from zoospore release from cul-tured aplanospores [prezoosporangia]). Tissuepreparation involves fixing concentratedzoospores or zoosporangia (produced by plac-ing the aplanospores into filtered ambient sea-water, where they develop into zoosporangia andproduce hundreds of motile zoospores) in 2-3%glutaraldehyde mixed and buffered for ambientfiltered seawater. Oyster tissues can also be fixedin 1G4F for 12-24 hrs. Following primary fixa-tion, rinse tissues in a suitable buffer and post-fix in 1-2% osmium tetroxide (OsO

4= osmic acid

- highly toxic). Secondary fixation should be com-plete within 1 hr. The OsO

4fixative must also be

rinsed with buffer/filtered (0.22 µm) seawaterprior to dehydration and resin-embedding.

Post-fixed tissues can be stored in a compatiblebuffer or embedded post-rinsing in a resin suit-able for ultramicrotome sectioning. Screening of1 ?m-thick sections melted onto glass micro-scope slides with 1% toluidine blue solution isone method of selecting the tissue specimensfor optimum evidence of putative Perkinsus spp.Such pre-screening should not be necessary forconcentrated zoospore or zoosporangia prepa-rations. Ultrathin sections are then mounted oncopper grids (with or without formvar coating),and stained with lead citrate + uranyl acetate (orequivalent EM stain).

The anterior flagellum of Perkinsus marinuszoospores is ornamented with a row of hair- andspur-like structures. The posterior flagellum issmooth. An apical complex is present, consist-ing of a conoid, sub-pellicular microtubules,rhoptries, rectilinear micronemes and conoid-associated micronemes. Large vacuoles are alsopresent at the anterior end of the zoospore.


haematoxylin-eosin) enable detection ofPerkinsus spp.

Perkinsus marinus infections are usually sys-temic, with trophozoites occuring in the connec-tive tissue of all organs. Immature trophozoites(meronts, merozoites or aplanospores) measure2-3 µm in diameter. “Signet-ring” stages are ma-ture trophozoites, measuring 3-10 µm in diam-eter, each with a visible eccentric vacuole dis-placing the nucleus and cytoplasm peripher-ally (Fig.M.5.3.2.1a). The “rosette” stage(tomonts, sporangia or schizonts) measure 4-15 µm in diameter and can contain 2, 4, 8, 16or 32 developing trophozoites (Fig.M.5.3.2.1b).

Perkinsus olseni shows the same developmen-tal stages although the trophozoite stages arelarger ranging from 13-16 µm in diameter. Dueto host and parasite diversity, however, mor-phological features cannot be considered spe-cific.

M.5.3.2.2 Fluid Thioglycollate Culture(Level II)

Tissue samples measuring 5-10 mm are excised(select lesions, rectal and gill tissues) and placedin fluid thioglycollate medium containing antibiot-ics. Incubation temperature and time varies perhost species and environment. The standard pro-tocol for P. marinus is 22-25˚C for 4-7 days inthe dark. Warmer temperatures can be used forP. olseni.

The cultured parasites expand in size to 70-250?m in diameter. Following incubation, the tissuesand placed in a solution of 1:5 Lugol’s iodine:waterfor 10 minutes. The tissue is then teased aparton a microscope slide and examined, using lowpower on a light microscope, for enlargedhypnospores with walls that stain blue-black(Fig.M.5.3.2.2).


More detailed methods for diagnostics can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.

M.5.4.1 Presumptive


Histology (M.5.3.2.1), may be used. However,for first-time diagnoses a back up tissue speci-men fixed for EM is recommended (M.5.4.2.1).

136


Proliferation of Perkinsus spp. is correlated withwarm water temperatures (>20˚C) and this co-incides with increased clinical signs and mor-talities. Effects appear cumulative with mortali-ties peaking at the end of the warm water sea-son in each hemisphere. The infective stage isa biflagellate zoospore which transforms intothe feeding trophozoite stage after entering thehost’s tissues. These multiply by binary fissionwithin the host tissues. Perkinsus marinusshows a wide salinity tolerance range. Perkinsusolseni is associated with full strength salinityenvironments.

Direct transmission of Perkinsus spp. has beendemonstrated by exposure of susceptible hoststo infected hosts, including cross-species trans-mission for P. olseni. There is currently no evi-dence of cross-genus transmission of P. marinus.


None known for Perkinsus spp. Most effortsagainst P. marinus have concentrated on devel-opment of resistant (tolerant) stocks of oysters.These currently show potential for surviving inenzootic areas, but are not recommended for usein non-enzootic areas due to their potential to actas sub-clinical carriers of the pathogen. Somesuccess has been achieved, however, in prevent-ing P. marinus infection of hatchery-reared larvaland juvenile oysters using filtration and UV ster-ilization of influent water. The almost ubiquitousoccurrence of Perkinsus in many bivalve spe-cies around mainland Australia makes control byrestriction of movements impractical.


Alemida, M., F.C. Berthe, A. Thebault, and M.T.Dinis. 1999. Whole clam culture as a quantita-tive diagnostic procedure of Perkinsusatlanticus (Apicomplexa, Perkinsea) in clamsRuditapes decussatus. Aquac. 177(1-4): 325-332.

Blackbourn, J., S.M. Bower, and G.R. Meyer.1998. Perkinsus qugwadi sp. nov. (incertaecedis), a pathogenic protozoan parasite ofJapanese scallops, Patinopecten yesssoensis,cultured in British Columbia, Canada. Can. J.Zool. 76(5): 942-953.

Bower, S.M., J. Blackbourn, and G.R. Meyer.1998. Distribution, prevalence and patho-genicity of the protozoan Perkinsus qugwadi

in Japanese scallops, Patinopectenyesssoensis, cultured in British Columbia,Canada. Can. J. Zool. 76(5): 954-959.

Bower, S.M., J. Blackbourne, G.R. Meyer, andD.W. Welch. 1999. Effect of Perkinsusqugwadi on various species and strains ofscallops. Dis. Aquat. Org. 36(2): 143-151.

Canestri-Trotti, G., E.M. Baccarani, F. Paesanti,and E. Turolla. 2000. Monitoring of infectionsby Protozoa of the genera Nematopsis,Perkinsus and Porospora in the smooth venusclam Callista chione from the north-westernAdriatic Sea (Italy). Dis. Aquat. Org. 42(2):157-161.

Cook, T., M. Folli, J. Klinck, S.E. Ford, and J.Miller. 1998. The relationship between increas-ing sea-surface temperature and the northwardspread of Perkinsus marinus (Dermo) diseaseepizootics in oysters. Estuarine, Coastal andShelf Science 46(4): 587 -597.

Fisher, W.S., L.M. Oliver, L., W.W. Walker, C.S.Manning and T.F. Lytle. 1999. Decreased re-sistance eastern oysters (Crassostreavirginica) to a protozoan pathogen (Perkinsusmarinus) after sub-lethal exposure to tributyltinoxide. Mar. Environ. Res. 47(2): 185-201.

Ford, S.E., R. Smolowitz, and M.M. Chintala.2000. Temperature and range extension byPerkinsus marinus. J. Shellfish Res. 19(1): 598(abstract).

Ford, S.E., Z. Xu, and G. Debrosse. 2001. Useof particle filtration and UV radiation to preventinfection by Haplosporidium nelsoni (MSX) andPerkinsus marinus (Dermo). Aquac. 194(1-2):37-49.

Hine, P.M. and T. Thorne. 2000. A survey of someparasites and diseases of several species ofbivalve mollusc in northern Western Austra-lia. Dis. Aquat. Org. 40(1): 67 -78.

Kotob, S.I., S.M. McLaughlin, P. van Berkum,and M. Faisal. 1999. Discrimination betweentwo Perkinsus spp. isolated from the soft shellclam, Mya arenaria, by sequence analysis oftwo internal transcribed spacer regions and the5.8S ribosomal RNA gene. Parasitol. 119(4):363-368.

Kotob, S.I., S.M. McLaughlin, P. van Berkum, P.and M. Faisal. 1999. Characterisation of twoPerkinsus spp. from the soft shell clam, Mya


137

arenaria, using the small subunit ribosomalRNA gene. J. Eukaryotic Microbiol. 46(4):439-444.

McLaughlin, S.M. and M. Faisal. 1998. In vitropropagation of two Perkinsus spp. from thesoft shell clam Mya arenaria. Parasite 5(4):341-348.

McLaughlin, S.M. and M. Faisal. 1998. Histo-pathological alternations associated withPerkinsus spp. infection in the soft shell clamMya arenaria. Parasite 5(4): 263-271.

McLaughlin, S.M. and M. Faisal. 1999. A com-parison of diagnostic assays for detection ofPerkinsus spp. in the soft shell clam Myaarenaria. Aquac. 172(1-2): 197-204.

McLaughlin, S.M. and M. Faisal. 2000. Preva-lence of Perkinsus spp. in Chesapeake Baysoft-shell clams, Mya arenaria Linnaeus, 1758,during 1990-1998. J. Shellfish Res. 19(1): 349-352.

Nickens, A.D., E. Wagner, and J.F. LaPeyre.2000. Improved procedure to count Perkinsusmarinus in eastern oyster hemolymph. J.Shellfish Res. 19(1): 665 (abstract).

O’Farrell, C.L., J.F. LaPeyre, K.T. Paynter, andE.M. Burreson. 2000. Osmotic tolerance andvolume regulation in in vitro cultures of theoyster pathogen Perkinsus marinus. J. Shell-fish Res. 19(1): 139-145.




Ordas, M.C. and A. Figueras. 1998. In vitro cul-ture of Perkinsus atlanticus, a parasite of thecarpet shell clam Ruditapes decussatus. Dis.Aquat. Org. 33(2): 129-136.

Ordas, M., A. Ordas, C. Beloso, and A. Figueras.2000. Immune parameters in carpet shell

clams naturally infected with Perkinsusatlanticus. Fish and Shellfish Immunol. 10(7):597-609.

Park, K-I., K-S. Choi, and J-W. Choi. 1999. Epi-zootiology of Perkinsus sp. found in Manilaclam, Ruditapes philippinarum in KomsoeBay, Korea. J. Kor. Fish. Soc. 32(3): 303-309.

Robledo, J.A.F., J.D. Gauthier, C.A. Coss, A.C,Wright, G.R. Vasta. 1999. Species -specific-ity and sensitivity of a PCR-based assay forPerkinsus marinus in the eastern oyster,Crassostrea virginica: A comparison with thefluid thioglycollate assay. J. Parasitol. 84(6):1237-1244.

Robledo, J.A.F., C.A. Coss, and G.R. Vasta.2000. Characterization of the ribosomal RNAlocus of Perkinsus atlanticus and developmentof a polymerase chain reaction -based diag-nostic assay. J. Parasitol. 86(5): 972-978.

Yarnall, H.A., K.S. Reece, N.A. Stokes, and E.M.Burreson. 2000. A quantitative competitivepolymerase chain reaction assay for the oys-ter pathogen Perkinsus marinus. J. Parasitol.86(4): 827-837.


138

M.6 HAPLOSPORIDIOSIS(HAPLOSPORIDIUM COSTALE,

H. NELSONI)

(PM Hine)

Fig.M.6.1.3b. Oil immersion magnification ofthe operculated spore stage of theHaplosporidium-like parasite in the gold-lippedpearl oyster Pinctada maxima from north West-ern Australia. Scale bar 10 µm. (H&E).

(PM Hine)

Fig.M.6.1.3c. Haemocyte infiltration activity inthe connective tissue of a Sydney rock oyster(Saccostrea cucullata) containing spores of aHaplosporidium-like parasite (arrow). Scale bar0.5 mm. (H&E).

(PM Hine)

Fig.M.6.1.3d. Oil immersion magnification ofHaplosporidium-like spores (arrow) associatedwith heavy haemocyte infiltration in a Sydneyrock oyster (Saccostrea cucullata). Scale bar10 µm. (H&E).

(PM Hine)

Fig.M.6.1.3a. Massive connective tissue anddigestive tubule infection by an unidentifiedHaplosporidium-like parasite in the gold-lippedpearl oyster Pinctada maxima from north West-ern Australia. Scale bar 0.5 mm (H&E).



Haplosporidiosis is caused by two species ofprotistan parasite belonging to the phylumHaplosporidia. Haplosporidium nelsoni (syn.Minchinia nelsoni) is responsible for “MSX” (multi-nucleate sphere X) disease in Crassostreavirginica (American oysters) and Haplosporidiumcostale (Minchinia costale) causes “SSO” (sea-side organism) disease in the same species.More information about the disease can be foundin the OIE Diagnostic Manual for Aquatic AnimalDiseases (OIE 2000a).

M.6.1.2 Host Range

Both Haplosporidium nelsoni and H. costalecause disease in Crassostrea virginica (Ameri-can oysters). Recently a Haplosporidium sp. fromCrassostrea gigas (Pacific oyster) has been iden-tified as H. nelsoni using DNA sequencing ofsmall sub-unit ribosomal DNA.


Haplosporidium nelsoni occurs in American oys-ters along the Atlantic coast of the United Statesfrom northern Florida to Maine. Enzootic areasappear limited to Delaware Bay, ChesapeakeBay, Long Island Sound and Cape Cod.Haplosporidium nelsoni has been found in C. gi-gas from California and Washington on the Pa-cific coast of the USA, and Korea, Japan andFrance.

139

M.6 Haplosporidiosis(Haplosporidium costale, H. nelsoni)

Haplosporidium costale has been reportedsolely from C. virginica from the Atlantic coastof the United States and has a small subunitrDNA distinct from that of H. nelsoni.Hapolosporidium costale also has a narrowerdistribution, ranging from Long Island Sound,New York, to Cape Charles, Virginia.

Similar agents have been reported from hatch-ery-reared pearl oysters, Pinctada maxima,(Fig.M.6.1.3a,b) and the rock oyster, Saccostreacucullata (Fig.M.6.1.3c,d), from north WesternAustralia.


No information or no positive report for this dis-ease in any country for the reporting periods 1999and 2000 (OIE 1999, OIE 2000b).


Haplosporidium nelsoni occurs extracellularlyin the connective tissue and digestive gland epi-thelia. It is often associated with a visible brown-red discolouration of gill and mantle tissues.Sporulation of H. nelsoni is prevalent in juvenileoysters (1-2 yrs) but sporadic in adults and oc-curs exclusively in the epithelial tissues of thedigestive tubules. Sporulation of H. costale oc-curs throughout the connective tissues.

Haplosporidum nelsoni infections appear andcontinue throughout the summer (mid-May to theend of October). Gradual disruption of the diges-tive gland epithelia is associated with weaken-ing and cumulative mortalities of oysters. A sec-ond wave of mortalities may occur in early springfrom oysters too weak to survive over-wintering.Holding in vivo for up to 2 weeks in 10 ppt salinityseawater at 20˚C kills the parasite but not thehost. H. nelsoni does not cause disease at <15ppt salinity.

Haplosporidium costale causes a pronouncedseasonal mortality between May and June.Sporulation is more synchronous than with MSXinfections, causing acute tissue disruption, weak-ening and death of heavily infected individuals.SSO disease is restricted to salinities of 25-33ppt and infections appear to be lost at lowersalinities.


More detailed methods for screening can befound in the OIE Diagnostic Manual for Aquatic

Animal Diseases (OIE 2000a), at http://www.oie.int or selected references.

M.6.3.1 Presumptive


Slowed growth, gaping and mortalities ofCrassostrea virginica and C. gigas should beconsidered suspect for Haplosporidiosis. Grosssigns are not pathogen specific and require LevelII examination, at least for first time observa-tions.


As with histology (M.6.3.2.2), juvenile oystersare preferred for cytological or tissue imprintscreening for Haplosporidium nelsoni. For H.costale adult oysters are preferred. Screeningduring May-June is recommended for both dis-ease agents.

Heart smears or impressions (dabs) can be madeonto a clean microscope slide and air-dried. Di-gestive gland and gill sections can also be usedfor smears by absorbing excess water from cutsurfaces and dabbing the surface onto cleanslides. Once dry, the slide is fixed in 70% metha-nol. Quick and effective staining can be achievedwith commercially available blood-staining (cy-tological) kits, using the manufacturer’s instruc-tions. The stained slides are then rinsed (gently)under tap water, allowed to dry, and cover-slippedusing a synthetic resin mounting medium.

The presence (especially between March andJune in endemic areas) of multinucleate plasmo-dia measuring 2-15 µm in diameter is indica-tive of H. costale infection (Fig.6.3.1.2a). Plas-modia of H. nelsoni are detectable betweenmid-May and October throughout the tissuesand measure 4-30 µm in diameter(Fig.M.6.3.1.2b).

Haemolymph suspensions can also be col-lected from live oysters, however, this is moretime-consuming than heart/tissue imprints andis considered less useful for screening pur-poses.


For Haplosporidium nelsoni, juvenile oysters arepreferred for screening. For H. costale adult oys-ters are preferred. Screening during May-Juneis recommended for both disease agents. The

140


(SE McGladdery)

Fig.M.6.4.2.2b. Oil immersion magnification ofMSX spores in the digestive tubule epitheliumof an American oyster Crassostrea virginica.Scale bar 25 µm. (H&E).

techniques used are the same as described forconfirmatory diagnosis (M.6.4.2.2).


More detailed methods for diagnostics can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.

M.6.4.1 Presumptive


The only presumptive diagnosis would be grossobservations of cumulative mortalities of Ameri-can oysters in early spring and late summer inareas (12-25 ppt salinity) with an established his-tory of MSX epizootics. Such presumptive diag-nosis requires confirmation via another diagnos-tic technique (histology). Likewise, summermortalities of the same oyster species in waterswith a history of SSO disease may be presumedto be due to SSO. Both must be confirmed, how-ever, since infection distributions for bothspecies of Haplosporidium may overlap.



Positive cytological or tissue imprints (M.6.4.1.1)can be considered confirmatory where collectedfrom susceptible oyster species and areas witha historic record of the presence ofHaplosporidium spp. infections.

(SE McGladdery)

Fig.M.6.3.1.2a. Plasmodia (black arrows) andspores (white arrows) of Haplosporidiumcostale, the cause of SSO disease, throughoutthe connective tissue of an American oyster(Crassostrea virginica). Scale bar 50 µm.

(SE McGladdery)

Fig.M.6.3.1.2b. Plasmodia (black arrows) andspores (white arrows) of Haplosporidiumnelsoni, the cause of MSX disease, throughoutthe connective tissue and digestive tubules ofan American oyster (Crassostrea virginica).Scale bar 100 µm.

(SE McGladdery)

Fig.M.6.4.2.2a. Oil immersion magnification ofSSO spores in the connective tissue of anAmerican oyster Crassostrea virginica. Scalebar 15 µm.

141


Positive histological sections can be consid-ered confirmatory where collected from suscep-tible oyster species and areas with a historicrecord of the presence of Haplosporidium spp.infections.

It is recommended that at least two dorso-ven-tral sections through each oyster be examinedusing oil immersion for screening purposes. Sec-tions from oysters >2 yrs (or >30 mm shellheight) should be fixed immediately in a fastfixative such as 1G4F. Davidson’s or 10% buff-ered formalin may be used for smaller or wholeoysters (see M.1.3.3.3) but these fixatives arenot optimal for subsequent confirmatory Elec-tron Microscopy (EM) diagnosis (M.6.4.2.3), orspecies identification, if required. Several stan-dard stains (e.g., haemotoxylin-eosin) enabledetection of Haplosprodium spp..

Haplosporidium spp. infections are usually sys-temic and characterised by massive infiltrationby hyalinocyte haemocytes (agranularhaemocytes with a low cytoplasm: nucleoplasmratio). The sporoplasm of spores of H. costalewhich are smaller than those of MSX and oftenmasked by the intense haemocyte infiltration re-sponse, can be differentially stained using amodified Ziehl-Nielsen stain. Sporocyts of H.costale occur in the connective tissues(Fig.M.6.3.1.2a), measure approximately 10-25µm in diameter and contain oval, operculate,spores approximately 3 µm in size(Fig.M.6.4.2.2a). Sporocysts of H. nelsoni oc-cur in the digestive tubule epithelia and mea-sure 20-50 µm in diameter. The operculatespores of MSX measure 4-6 x 5-8 µm(Fig.M.6.4.2.2b). In C. gigas spores may alsooccur in other tissues. Older foci of infection inboth oyster species may be surrounded byhaemocytes and necrotic tissue debris. A simi-lar infectious agent occurs in the pearl oysterPinctada maxima in north Western Australia(Fig.M.6.1.3a,b). The spore size of thisHaplosporidium resembles H. nelsoni but dif-fers from MSX infections in both C. virginicaand C. gigas by being found exclusively in theconnective tissue.

The plasmodial stages of both H. costale and H.nelsoni are as described under M.6.3.1.2.

1 Attention Dr. N. Stokes, Virginia Institute of Marine Science, College of William and Mary, Gloucester Point, Virginia 23062,USA. (E-mail: [email protected]).



TEM is required for confirmation of species-spe-cific ultrastructure of the spores - especially inareas enzootic for both disease agents. Tissuesare fixed in 2-3% glutaraldehyde mixed andbuffered for ambient filtered seawater. Oystertissues can also be fixed in 1G4F for 12-24 hrs.Following primary fixation, rinse tissues in asuitable buffer and post-fix in 1-2% osmiumtetroxide (OsO4 = osmic acid - highly toxic).Secondary fixation should be complete within1 hr. The OsO4 fixative must also be rinsed withbuffer/filtered (0.22 µm) seawater prior to de-hydration and resin-embedding.Post-fixed tissues can be stored in a compat-ible buffer or embedded post-rinsing in a resinsuitable for ultramicrotome sectioning. Screen-ing of 1 micron sections melted onto glassmicroscope slides with 1% toluidine blue solu-tion is one method of selecting the tissue speci-mens for optimum evidence of Haplosporidiumplasmodia or spores.

M.6.4.2.4 In situ Hybridization (Level III)

DNA-probes for both species of Haplosporidiumhave been produced at the Virginia Institute ofMarine Science (VIMS), College of William andMary, Gloucester, Virginia, USA. These are notyet commercially available, but labelled probesmay be obtained for experienced users, orsamples may be sent to VIMS1 for in situ hy-bridization analysis.


Neither parasite has been successfully transmit-ted under laboratory conditions and one (or more)intermediate host(s) is/are suspected.


None are known for Haplosporidium spp.. Mostefforts have concentrated on development of re-sistant stocks of oysters. These currently showpotential for survival in enzootic areas, but arenot recommended for use in non-enzootic areasdue to their potential as sub-clinical carriers ofthe pathogen. Some success has also beenachieved in preventing infection of hatchery-reared larval and juvenile oysters through filtra-tion and UV radiation of influent water.

142


Andrews, J.D. 1967. Interaction of two diseasesof oysters in natural waters. Proc. Nat. Shell-fish. Assoc. 57: 38-49.

Andrews, J.D. 1982. Epizootiology of late sum-mer and fall infections of oysters byHaplosporidium nelsoni, and comparison to theannual life cycle of Haplosporidium costalis, atypical haplosporidian. J. Shellfish Res. 2: 15-23.

Andrews, J.D. and M. Castagna. 1978. Epizooti-ology of Minchinia costalis in susceptibleoysters in seaside bays of Virginia’s easternshore, 1959-1976. J. Inverteb. Pathol. 32:124-138.

Andrews, J.D., J.L. Wood, and H.D. Hoese.1962. Oyster mortality studies in Virginia: III.Epizootiology of a disease caused byHaplosporidium costale, Wood and Andrews.J. Insect Pathol. 4(3): 327-343.

Barber, B.J., S.E. Ford, and D.T.J. Littlewood.1991. A physiological comparison of resistantand susceptible oysters Crassostrea virginica(Gmelin) exposed to the endoparasiteHaplosporidium nelsoni (Haskin, Stauber &Mackin). J. Exper. Mar. Biol. Ecol. 146: 101-112.

Burreson, E.M. 1997. Molecular evidence for anexotic pathogen: Pacific origin ofHalposporidium nelsoni (MSX), a pathogen ofAtlantic oysters, p. 62. In: M. Pascoe (ed.). 10thInternational Congress of Protozoology, TheUniversity of Sydney, Australia, Monday 21July - Friday 25 July 1997, Programme & Ab-stracts. Business Meetings & Incentives,Sydney. (abstract).

Burreson, E.M., M.E. Robinson, and A. Villalba.1988. A comparison of paraffin histology andhemolymph analysis for the diagnosis ofHaplosporidium nelsoni (MSX) in Crassostreavirginica (Gmelin). J. Shellfish Res. 7: 19-23.

Burreson, E.M., N.A. Stokes, and C.S. Friedman.2000. Increased virulence in an introducedpathogen: Haplosporidium nelsoni (MSX) in theeastern oyster Crassostrea virginica. J. Aquat.Anim. Health 12(1): 1-8.

Comps, M. and Y. Pichot. 1991. Fine spore struc-ture of a haplosporidan parasitizingCrassostrea gigas: taxonomic implications.Dis. Aquat. Org. 11: 73-77.

Farley, C.A. 1967. A proposed life-cycle ofMinchinia nelsoni (Haplosporida,Haplosporidiidae) in the American oysterCrassostrea virginica. J. Protozool. 22(3):418-427.

Friedman, C.S., D.F. Cloney, D. Manzer, andR.P. Hedrick. 1991. Haplosporidiosis of thePacific oyster, Crassostrea gigas. J. Inverteb.Pathol. 58: 367-372.

Fong, D., M.-Y. Chan, R. Rodriguez, C.-C. Chen,Y. Liang, D.T.J. Littlewood, and S.E. Ford.1993. Small subunit ribosomal RNA gene se-quence of the parasitic protozoanHaplosporidium nelsoni provides a molecu-lar probe for the oyster MSX disease. Mol.Biochem. Parasitol. 62: 139-142.

Ford, S.E. 1985. Effects of salinity on survivalof the MSX parasite Haplosporidium nelsoni(Haskin, Stauber and Mackin) in oysters. J.Shellfish Res. 5(2): 85-90.

Ford, S.E. 1992. Avoiding the transmission ofdisease in commercial culture of molluscs, withspecial reference to Perkinsus marinus(Dermo) and Haplosporidium nelsoni (MSX).J. Shellfish Res. 11: 539-546.

Ford, S.E. and H.H. Haskin. 1987. Infection andmortality patterns in strains of oystersCrassostrea virginica selected for resistanceto the parasite Haplosporidium nelsoni (MSX).J. Protozool. 73(2): 368-376.

Ford, S.E. and H.H. Haskin. 1988. Managementstrategies for MSX (Haplosporidium nelsoni)disease in eastern oysters. Amer. Fish. Soc.Spec. Pub. 18: 249?256.

Ford, S.E., Z. Xu, and G. Debrosse. 2001. Useof particle filtration and UV radiation to preventinfection by Haplosporidium nelsoni (MSX) andPerkinsus marinus (Dermo) in hatchery-rearedlarval and juvenile oysters. Aquac. 194(1-2):37-49.

Hine, P.M. and T. Thorne. 2000. A survey of someparasites and diseases of several species ofbivalve mollusc in northern Western Austra-lia. Dis. Aquat. Org. 40(1): 67-78.

Katkansky, S.C. and R.W. Warner. 1970. Sporu-lation of a haplosporidian in a Pacific oyster(Crassostrea gigas) in Humboldt Bay, Cali-fornia. J. Fish. Res. Bd. Can. 27(7): 1320-1321.


143


Kern, F.G. 1976. Sporulation of Minchinia sp.(Haplosporida, Haplosporidiidae) in the Pa-cific oyster Crassostrea gigas (Thunberg) fromthe Republic of Korea. J. Protozool. 23(4):498-500.




Renault, T., N.A. Stokes, B. Chollet, N.Cochennec, F.C. Berthe, A. Gerard, A. andE.M. Burreson. 2000. Haplosporidiosis in thePacific oyster Crassostrea gigas from theFrench Atlantic coast. Dis. Aquat. Org. 42(3):207-214.

Stokes, N.A. and E.M. Burreson. 1995. A sen-sitive and specific DNA probe for the oysterpathogen Haplosporidium nelsoni. J. Eukary-otic Microbiol. 42: 350-357.

Wolf, P.H. and V. Spague. 1978. An unidentifiedprotistan parasite of the pearl oyster Pinctadamaxima, in tropical Australia. J. Inverteb.Pathol. 31: 262-263.

144



Marteilioidosis is caused by two species of para-sites, belonging to the protistan PhylumParamyxea. Marteilioides chungmuenis is re-sponsible for oocyte infections of Pacific oysters(Crassostrea gigas) and Marteilioides branchialisinfects the gills of Saccostrea glomerata (syn.Crassostrea commercialis, Saccostreacommercialis).

M.7.1.2 Host Range

The Pacific oyster Crassostrea gigas is infectedby Marteilioides chungmuensis. Marteilioidesbranchialis infects the Sydney rock oyster,Saccostrea commercialis.


Marteilioides chungmuensis infects C. gigas inJapan and Korea. Marteilioides branchialis isfound in Australia (New South Wales).


Marteilioides chungmuensis infects the cytoplasmof mature oocytes and significant proportions ofthe reproductive output of a female oyster canbe affected. The infected eggs are released orretained within the follicle, leading to visible dis-tention of the mantle surface (Fig.M.7.2a, b).Prevalences of up to 8.3% have been reportedfrom Korea. Marteilioides branchialis causes fo-cal lesions on the gill lamellae and, in conjunc-tion with infections by Marteilia sydneyi (M.3), isassociated with significant mortalities of Sydneyrock oysters being cultured in trays during theautumn.


M.7.3.1 Presumptive


Marteilioides branchialis causes focal patches (1-2 mm in diameter) of discolouration and swellingon the gill lamellae. The presence of such lesionsin Sydney rock oysters in the Austral autumnshould be treated as presumptive Marteilioidosis.



M.7 MARTEILIOIDOSIS(MARTEILIOIDES CHUNGMUENSIS,

M. BRANCHIALIS)

The techniques used are the same as describedfor confirmatory disease diagnosis (M.7.4.2.1).Presence of histological inclusions, as describedunder M.7.4.2.1, can be considered confirmatoryfor Marteilioides spp., during screening.

(MS Park and DL Choi)

Fig.M.7.2a,b. a. Gross deformation of mantletissues of Pacific oyster (Crassostrea gigas)from Korea, due to infection by the protistanparasite Marteiloides chungmuensis causingretention of the infected ova within the ovaryand gonoducts; b. (insert) normal mantle tis-sues of a Pacific oyster.

(MS Park)

Fig.M.7.4.2.1. Histological section through theovary of a Pacific oyster (Crassostrea gigas) withnormal ova (white arrows) and ova severely in-fected by the protistan parasite Marteiliodeschungmuensis (black arrows). Scale bar 100µm.


M.7.4.1 Presumptive


As for M.7.3.1.1, focal patches (1-2 mm in diam-eter) of discolouration and swelling on the gilllamellae of Sydney rock oysters in the Australautumn can be treated as presumptive positivesfor M. branchialis.

145


For first-time diagnoses a back up tissue speci-men fixed for EM is recommended (M.7.4.2.3).



Positive histological sections can be consideredconfirmatory where collected from susceptibleoyster species and areas with a historic recordof the presence of Marteilioides spp. infections.

It is recommended that at least two dorso-ven-tral sections through each oyster be examinedusing oil immersion for screening purposes. Sec-tions from oysters >2 yrs (or >30 mm shell height)should be fixed immediately in a fast fixative suchas 1G4F. Davidsons or 10% buffered formalinmay be used for smaller or whole oysters (seeM.1.3.3.3) but these fixatives are not optimal forsubsequent confirmatory Electron Microscopy(EM) diagnosis (M.6.4.2.3), or species identifi-cation, if required. Several standard stains (e.g.,haemotoxylin-eosin) enable detection ofMarteilioides spp..

Marteilioides chungmuensis is located in the cy-toplasm of infected ova (Fig.M.7.4.2.1). Stem(primary) cells contain secondary cells. Thesemay, in turn, contain developing sporonts, givingrise to a single tertiary cell by endogenous bud-ding. Each tertiary cell forms a tricellular sporeby internal cleavage.

Marteilioides branchialis causes epithelial hyper-plasia and granulocyte infiltration at the site ofinfection. Uninucleate primary cells contain twoto six secondary cells (some may contain up to12) in the cytoplasm of epithelial cells, connec-tive tissue cells and occasionally the infiltratinghaemocytes within the lesion.


TEM is required for confirmation of species-spe-cific ultrastructure of these parasites. Tissues arefixed in 2-3% glutaraldehyde mixed and bufferedfor ambient filtered seawater. Tissues can alsobe fixed in 1G4F for 12-24 hours. Following pri-mary fixation, rinse in a suitable buffer and post-fix in 1-2% osmium tetroxide (OsO

4= osmic acid

- highly toxic). Secondary fixation should be com-plete within 1 hour. The OsO

4fixative must also

be rinsed with buffer/filtered (0.22 microns) sea-water prior to dehydration and resin-embed-ding.

M.7 Marteilioidosis(Marteilioides chungmuensis,

M. branchialis)

Post-fixed tissues should be stored in a com-patible buffer or embedded post-rinsing in aresin suitable for ultramicrotome sectioning.Screening of 1 micron sections melted ontoglass microscope slides with 1% toludine bluesolution is one method of selecting the tissuespecimens for optimum evidence of putativeMarteilioides spp. Ultrathin sections are thenmounted on copper grids (with or withoutformvar reinforced support) for staining withlead citrate + uranyl acetate or equivalent EMstain.

Marteilioides branchialis is differentiated from theother Marteilioides spp. by the presence of twoconcentric cells (rather than three) within thespore. In addition M. chungmuensis in C. gigascontains only two to three sporonts per primary/stem cell compared with two-six (or up to 12) forM. branchialis. Multivesicular bodies resemblingthose of Marteilia spp. are present in M.branchialis stem cells, but absent from those ofM. chungmuensis.


Unknown.


None known.


Anderson, T.J. and R.J.G. Lester. 1992.Sporulation of Marteilioides branchialis n.sp.(Paramyxea) in the Sydney rock oyster,Saccostrea commercialis: an electronmicroscope study. J. Protozool. 39(4): 502-508.

Anderson, T.J., T.F. McCaul, V. Boulo, J.A.F.Robledo, and R.J.G. Lester. 1994. Light andelectron immunohistochemical assays onparamyxea parasites. Aquat. Living Res. 7(1):47-52.

Elston, R.A. 1993. Infectious diseases of thePacific oyster Crassostrea gigas. Ann. Rev.Fish Dis. 3: 259-276.

Comps, M., M.S. Park, and I. Desportes. 1986.Etude ultrastructurale de Marteilioideschungmuensis n.g. n.sp., parasite desovocytes de l’hu?tre Crassostrea gigas Th.Protistol. 22(3): 279-285.

146

M.7 Marteilioidosis(Marteilioides chungmuensis,

M. branchialis)

Comps, M., M.S. Park, and I. Desportes. 1987.Fine structure of Marteilioides chungmuensisn.g. n.sp., parasite of the oocytes of theoyster Crassostrea gigas. Aquac. 67(1-2):264-265.

Hine, P.M. and T. Thorne. 2000. A survey ofsome parasites and diseases of severalspecies of bivalve mollusc in northernWestern Australia. Dis. Aquat. Org. 40(1): 67-8.

147



Oyster Velar Virus Disease (OVVD) (Iridovirosis)is caused by an icosahedral DNA virus with mor-phological similarities to the Iridoviridae.

M.8.1.2 Host Range

Crassostrea gigas (Pacific oyster) larvae are thedocumented host species, although similar viralagents have been associated with gill disease(“Maladie des Branchies”) and haemocyte infec-tions in Portugese oysters (Crassostrea angulata)and C. gigas.


Infections have been reported solely from twohatcheries in Washington State, but are believedto have a ubiquitious distribution throughout ju-venile C. gigas production, with clinical manifes-tation only under sub-optimal growing conditions.


OVVD causes sloughing of the velar epitheliumof larvae >150µm in length, and can cause upto 100% mortality under hatchery conditions.The larvae can not feed, weaken and die.


M.8.3.1 Presumptive

Generally-speaking, since this is an opportunis-tic infection, only clinical infections will demon-strate detectable infections – as described underM.8.4.

M.8.3.1.1 Wet Mounts (Level I)

Wet mounts of veliger larvae which demonstratesloughing of ciliated epithelial surfaces can beconsidered to be suspect for OVVD. As withgross observations, other opportunistic patho-gens (bacteria and Herpes-like viruses) may beinvolved, so Level II/III diagnostics are required.


Using the techniques described under M.8.4.2.1,detection the features described in that sectioncan be considered to be presumptively positivefor OVVD. Such inclusions require TEM (LevelIII) (M.8.4.2.2) for confirmatory diagnosis, at leastfor first time observations.

M.8 IRIDOVIROSIS(OYSTER VELAR VIRUS DISEASE)


M.8.3.2.1 Transmission Electron Micros-copy (Level III)

As described under M.8.4.2.2.


M.8.4.1 Presumptive


Slowed growth, cessation of feeding and swim-ming in larval Crassostrea gigas should be con-sidered suspect for OVVD. Gross signs are notpathogen specific and require Level II examina-tion (M.8.4.2), at least for first time observations.

M.8.4.1.2 Wet Mounts (Level I)

As described under M.8.3.1.1. For first-time di-agnoses a back up tissue specimen fixed for TEMis recommended (M.8.4.2.2).



M.8.4.1.4 Transmission Electron Micros-copy (Level III)




Where larvae have a history of OVVD, detectionof inclusions and ciliated epithelial pathology, asdescribed below, can be considered confirma-tory for the disease. However, it should be notedthat other microbial infections can induce similarhistopathology and electron microscopy is theideal confirmatory technique (M.8.4.2.2).

Larvae must be concentrated by centrifugationor filtration into a pellet prior to embedding. Thisis best achieved post-fixation in Davidson’s,1G4F or other fixative. Although paraffin embed-ding is possible, resin embedding is recom-mended for optimal sectioning. Paraffin permitssectioning down to 3 µm using standard mi-crotome. Resin embedded tissue can be sec-tioned down to 1 µm thick, but requiresspecialised microtomes and/or block holdersand specialised staining.

148

Standard stains (e.g., haemotoxylin-eosin) willdetect intracytoplasmic inclusion bodies in cili-ated velar epithelial cells. Early inclusion bodiesare spherical, but become more irregular as theviruses proliferate. Inclusion bodies may also bedetected in oesophageal and oral epithelia or,more rarely, in mantle epithelial cells.


TEM is required to visualise the causative virusesin situ in gill tissue sections of concentrated ‘pel-lets’ of larvae. Fixation in 2-3% glutaraldehydemixed and buffered for ambient filtered seawatershould not exceed 1 hour to reduce artifacts. Tis-sues can also be fixed in 1G4F for 12-24 hrs.Following primary fixation, rinse in a suitablebuffer and post-fix in 1-2% osmium tetroxide(OsO4 = osmic acid - highly toxic). Secondaryfixation should be complete within 1 hr. The OsO4

fixative must also be rinsed with buffer/filtered(0.22 µm) seawater prior to dehydration andresin-embedding.

Post-fixed tissues should be stored in a compat-ible buffer or embedded post-rinsing in a resinsuitable for ultramicrotome sectioning. Screen-ing of 1 micron sections melted onto glass mi-croscope slides with 1% toludine blue solution isone method of selecting the best specimens forultrathin sectioning. Ultrathin sections aremounted on copper grids (with or without formvarreinforced support) for staining with lead citrate+ uranyl acetate or equivalent EM stain.

Icosahedral viral particles (228 +/- 7 nm in diam-eter) with a bi-laminar membrane capsid shouldbe evident to confirm Iridoviral involvement.


The disease appeared in March-May at affectedhatcheries. Direct transmission between mori-bund and uninfected larvae is suspected.


None known except for reduced stocking densi-ties, improved water exchange and generalhatchery sanitation methods (tank and line dis-infection, etc.).


Comps, M. and N. Cochennec. 1993. A Herpes-like virus from the European oyster Ostreaedulis L. J. Inverteb. Pathol. 62: 201-203.

Elston, R.A. 1979. Virus-like particles associ-ated with lesions in larval Pacific oysters(Crassostrea gigas). J. Inverteb. Pathol. 33:71-74.

Elston, R.A. 1993. Infectious diseases of thePacific oyster, Crassostrea gigas. Ann. Rev.Fish Dis. 3: 259-276.

Elston, R. 1997. Special topic review: bivalvemollusc viruses. Wor. J. Microbiol. Biotech. 13:393-403.

Elston, R.A. and M.T. Wilkinson. 1985. Pathol-ogy, management and diagnosis of oyster ve-lar virus disease (OVVD). Aquac. 48: 189-210.

Farley, C.A. 1976. Epizootic neoplasia in bivalvemolluscs. Prog. Exper. Tumor Res. 20: 283-294.

Farley, C.A., W.G. Banfield, G. Kasnic Jr. andW.S. Foster. 1972. Oyster Herpes-type virus.Science 178: 759-760.

Hine, P.M., B. Wesney and B.E. Hay. 1992. Her-pes virus associated with mortalities amonghatchery-reared larval Pacific oystersCrassostrea gigas. Dis. Aquat. Org. 12: 135-142.

Le Deuff, R.M., J.L. Nicolas, T. Renault and N.Cochennec. 1994. Experimental transmissionof a Herpes-like virus to axenic larvae of Pa-cific oyster, Crassostrea gigas. Bull. Eur.Assoc. Fish Pathol.14: 69-72.

LeDeuff, R.M., T. Renault and A. G?rard. 1996.Effects of temperature on herpes-like virusdetection among hatchery-reared larval Pacificoyster Crassostrea gigas. Dis. Aquat. Org. 24:149-157.

Meyers, T.R. 1981. Endemic diseases of culturedshellfish of Long Island, New York: adult andjuvenile American oysters (Crassostreavirginica) and hard clams(Mercenariamercenaria). Aquac. 22: 305-330.

Nicolas, J.L., M. Comps and N. Cochennec.1992. Herpes-like virus infecting Pacific oys-ter larvae, Crassostrea gigas. Bull. Eur. Assoc.Fish Pathol. 12: 11-13.

Renault, T., N. Cochennec, R.M. Le Deuff andB. Chollet. 1994. Herpes-like virus infectingJapanese oyster (Crassostrea gigas) spat.Bull. Eur.Assoc. Fish Pathol. 14: 64 -66.

M.8 Iridovirosis(Oyster Velar Virus Disease)

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Disease Expert/Laboratory

Mollusc pathogens Dr. F. BerthIFREMERLaboratoire de Genetique Aquaculture et PathologieBP 133, 17390 La TrembladeFRANCETel: 33(0)5 46.36.98.36Fax: 33 (0)5 46.36.37.51E-mail: [email protected]

Annex M.AI. OIE Reference Laboratoryfor Molluscan Diseases

150

Disease Expert

Bonamiosis Dr. Brian JonesSenior Fish PathologistFisheries WAAdjunct Professor, Muresk Institutec/o Animal Health Labs3 Baron-Hay CourtSouth Perth WA 6151AUSTRALIATel: 61-8-9368-3649Fax: 61-8-9474-1881E-mail: [email protected]/MicrocytosisDr. Robert D. AdlardQueensland MuseumPO Box 3300, South Brisbane,Queensland 4101AUSTRALIATel: +61 7 38407723Fax: +61 7 38461226E-mail: [email protected];http://www.qmuseum.qld.gov.au

Marteiliosis Dr. Sarah N. KleemanAquatic Animal BiosecurityAnimal BiosecurityGPO Box 858Canberra ACT 2601AUSTRALIATel: +61 2 6272 3024Fax: +61 2 6272 3399E-mail: [email protected]

Molluscan Diseases Professor R.J.G. LesterDepartment of Microbiology and ParasitologyThe University of Queensland, BrisbaneAUSTRALIA 4072.Tel: +61-7-3365-3305,Fax: +61-7-3365-4620E-mail: [email protected]://www.biosci.uq.edu.au/micro/academic/lesterlester.htmDr. Dong Lim ChoiPathology Division #408-1, Shirang-ri, Kijang-upKijang-gun, Busan 619-902KOREA ROTel: +82-51-720-2493Fax: +82-51-720-2498E-mail: [email protected]. Mi-Seon ParkPathology Division#408-1, Shirang-ri, Kijang-upKijang-gun, Busan 619-902KOREA RO 619-902Tel: +82-51-720-2493Fax: +82-51-720-2498E-mail: [email protected]

Annex M.AII. List of Regional Resource Expertsfor Molluscan Diseases Asia-Pacific1

1 The experts included in this list has previously been consulted and agreed to provide valuable information and health adviseconcerning their particular expertise.

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Dr. Jie HuangYellow Sea Fisheries Research InstituteChinese Academy of Fishery Sciences106 Nanjing RoadQingdao, Shandong 266071PEOPLE’S REPUBLIC of CHINATel: 86 (532) 582 3062Fax: 86 (532) 581 1514E-mail: [email protected]. Arthur de VeraFish Health SectionBureau of Fisheries and Aquatic ResourcesArcadia Building, 860 Quezon AvenueQuezon City, Metro ManilaPHILIPPINESFax: (632) 3725055Tel: (632) 4109988 to 89E-mail: [email protected]. Paul Michael HineAquatic Animal DiseasesNational Centre for Disease InvestigationMAF Operations, P.O. Box 40-742Upper HuttNEW ZEALANDTel: +64-4-526-5600Fax: +64-4-526-5601E-mail: [email protected]

List of Experts Outside Asia-Pacific Region2

Molluscan Diseases Dr. Susan BowerDFO Pacific Biological Station3190 Hammond Bay RoadNanaimo, British ColumbiaV9R 5K6CANADATel: 250-756-7077Fax: 250-756-7053E-mail: [email protected]. Sharon E. McGladderyOceans and Aquaculture Science200 Kent Street (8W160)Ottawa, Ontario, K1A 0E6CANADATel: 613-991-6855Fax: 613-954-0807E-mail: [email protected]

2 These experts outside the Asia-Pacific region has supported the regional programme on aquatic animal health and agreed toassist further in providing valuable information and health advise on molluscan diseases.

Annex M.AII. List of Regional Resource Experts forMolluscan Diseases Asia-Pacific

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• Australian Aquatic Animal Disease – Identification Field Guide (1999) by Alistair Herfortand Grant RawlinInformation: AFFA Shopfront – Agriculture, Fisheries and Forestry – Australia

GPO Box 858, Canberra, ACT 2601Tel: (02) 6272 5550 or free call: 1800 020 157Fax: (02) 6272 5771E-mail: [email protected]

• Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish byBower, SE McGladdery and IM Price (1994)Information: Dr. Susan Bower

DFO Pacific Biological Station3190 Hammond Bay RoadNanaimo, British ColumbiaV9R 5K6CANADATel: 250-756-7077Fax: 250-756-7053E-mail: [email protected]

• Mollusc Diseases: Guide for the Shellfish Farmer. 1990. by R.A. Elston. Washington SeaGrant Program, University of Washington Press, Seattle. 73 pp.

• A Manual of the Parasites, Pests and Diseases of Canadian Atlantic Bivalves. 1993. bySE McGladdery, RE Drinnan and MF Stephenson.Information: Dr. Sharon McGladdery

Oceans and Aquaculture Science200 Kent Street (8W160)Ottawa, Ontario, K1A 0E6Tel: 613-991-6855Fax: 613-954-0807E-mail: [email protected]

Annex M.AIII. List of Useful Diagnostic Manuals/Guides/Keys to Molluscan Diseases

basic anatomy of an oyster

Documents

pteria penguin shell

shell surface observationsm

inner shell observationsm

gross observationsm

environmental observationsm

sample collection

hard shell clam

bondad reantasofig