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Histopathology in Pacific oyster (Crassostrea gigas) spat caused by the dinoflagellate Prorocentrum rhathymum
Harmful Algae 4 (2005) 61–74
Histopathology in Pacific oyster (Crassostrea gigas) spat caused by the dinoflagellate Prorocentrum rhathymum Imojen Pearce a , Judith H. Handlinger b , Gustaaf M. Hallegraeff a,∗ a
b
School of Plant Science, University of Tasmania, Private Bag 55, Hobart 7001, Tasmania, Australia Fish Health Unit, Animal Health Laboratory, Department of Primary Industries Water and the Environment, P.O. Box 46, Kings Meadows 7249, Tasmania, Australia Received 24 August 2003; received in revised form 15 October 2003; accepted 15 November 2003
Abstract The recognition of an apparent association between seasonal oyster spat mortalities (up to 40%) and high Prorocentrum rhathymum density in the Little Swanport Estuary, Tasmania, prompted further experimental investigation into the toxicity by this dinoflagellate. Standard brine shrimp, haemolysis assays and intraperitoneal mouse bioassays revealed fast acting toxins in methanol but not aqueous extracts of P. rhathymum, with mice dying in less than 20 min. Oyster bioassays involved feeding spat (4 mm shell width) for 21 consecutive days on a diet of cultured P. rhathymum at simulated bloom densities (104 cells ml−1 ). No oyster mortality was observed, however, histopathological signs of thin, dilated gut tubules and sloughing of gut cells resembled those seen in affected field samples. In contrast to field samples, gill pathology was also observed in experimental exposure oysters. © 2003 Elsevier B.V. All rights reserved. Keywords: Crassostrea gigas; Histopathology; Oyster; Prorocentrum; Tasmania
1. Introduction A successful hatchery based Pacific oyster (Crassostrea gigas) industry has been established in Tasmania, Australia since the early 1980s. Oyster spat are produced for sale to commercial farmers, initially in a land-based nursery system of flowing unfiltered seawater, and subsequently are transferred to outgrow areas on racks in estuaries. Little Swanport is such a site with leases situated throughout the estuary. The leases are often established over extensive seagrass (Zostera muelleri) beds, the leaf shoots of which support dense ∗ Corresponding author. Tel.: +61-3-62262623; fax: +61-3-62262698. E-mail address: [email protected] (G.M. Hallegraeff).
algal epiphyte growth, particularly in the proximity of the intertidal oyster racks (Rees, 1993). A significant episode of site specific losses of post-hatchery spat was first noted in early January 1993, involving stock from several hatcheries in one of two nursery bays shared by these hatcheries. Smaller similar losses have since occurred at the same site, with notable reoccurrences in May 1998, May–June 2000, May 2001 and March–May 2002. Losses varied between batches present on the site, with maximum losses over several weeks of up to 40% on some racks. Histological, bacteriological and epidemiological study of these outbreaks indicated that the cause was not infectious, and no association could be found with artificial pollutants or pesticides from equipment within the bay or from agricultural or other
1568-9883/$ – see front matter © 2003 Elsevier B.V. All rights reserved. doi:10.1016/j.hal.2003.11.002
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activities upstream (Elston, 1999). However, there was evidence that these losses were more common in low rainfall years, and often followed the first significant rains. The oyster deaths were considered unlikely to be the direct result of freshwater run-off, as spat are usually well able to tolerate reduced salinities, and deaths were not immediate following rains but persisted for up to several weeks, sometimes despite a lack of further rain. The possibility of an origin in soil run-off was also considered. However, while potential areas of acid sulphate soil did exist in the Little Swanport catchment, there was no evidence that these had been drained. Similarly, iron from soil erosion was discounted as no tissue evidence was found in damaged spat. Initial algal studies showed the dinoflagellates Ostreopsis siamensis Schmidt and species of Prorocentrum Ehrenberg were present during the 1998 mortality (Elston, 1999). The involvement of toxic algae was further investigated during 1999, as part of a survey of the potentially toxic epiphytic algal assemblage on the seagrass in the estuary (Pearce et al., 2001). This assemblage included the dinoflagellates O. siamensis, Coolia monotis Meunier, Prorocentrum rhathymum Loeblich, Sherley and Schmidt, and Prorocentrum lima (Ehrenberg) Dodge, strains of which have been shown elsewhere to produce toxins (Holmes et al., 1995; Usami et al., 1995; Yasumoto et al., 1987). Water samples examined during the following periods of high spat mortality (March 2000 and March 2001), revealed that P. rhathymum was blooming on both occasions. This species was initially reported as Prorocentrum mexicanum, but was redefined as P. rhathymum following the redefinition of the two species by Cortés-Altamirano and Sierra-Beltrán (2003). Blooms of P. rhathymum have not previously been reported as causing damage to shellfish. However, the species has been identified as toxic (Tindall et al., 1989; Nakajima et al., 1981) and shellfish mortality has been linked to other members of the genus. Prorocentrum minimum (Pavillard) Schiller has been found to cause death in juvenile oysters and scallops in laboratory feeding experiments (Luckenbach et al., 1993; Wikfors and Smolowitz, 1995), an unidentified Prorocentrum sp. caused mortality of hundreds of flat oysters in China (Zhang et al., 1995) and human poisonings (diarrhetic shellfish poisoning) have been commonly associated with okadaic acid producing
species such as P. lima, Prorocentrum hoffmannianum Faust and Prorocentrum concavum Fukuyo (Dickey et al., 1990; Faust et al., 1999; Morton et al., 1994). In the present work, we aimed to document the characteristics of the recurrent oyster mortality at this site and through an extensive investigation between May 2001 and May 2002, determined whether the Tasmanian strains of P. rhathymum were toxic and responsible for the observed pathology of oyster spat.
2. Methods 2.1. Characterisation of the natural mortality events 2.1.1. Site description Little Swanport is located at the basis of the estuary formed by the Little Swanport River on the western side of Great Oyster Bay, approximately 30 km south of Swansea, Tasmania (Fig. 1). It has a complex hydrography due to its narrow entrance, long channel, numerous shoals and the irregular shape of the estuary. Water circulation is complex, being largely wind driven and is well mixed, as validated by temperature and salinity profiles (DPIWE, 1998). The flushing time is 2.3 tidal cycles or just over a day. Occasionally flooding of the Little Swanport River causes inundation of freshwater into the estuary, however, rainfall rarely exceeds 800 mm per annum (DPIWE, 1998). The surrounding land use is predominantly rural with some residential properties on the northern side of the estuary. 2.1.2. Background: histopathology and health monitoring The health of oysters in the Little Swanport Estuary has been monitored over the past decade through routine health surveillance as part of the Tasmanian Oyster Health Program, and through a Co-operative Research Centre for Aquaculture Project on early mollusc mortality in Tasmania. Histopathology was examined from farm spat undergoing mortality in Little Swanport on archived samples from the early 1993 losses, and samples from mortality episodes in May–June 1998, May 2000 and May 2002. These were compared with routine
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Fig. 1. Study site, Little Swanport, Tasmania (scale bar = 1 km): (1) nursery site (NJ); (2) lease L1–L2; (3) lease L7–L8.
6-monthly surveillance spat samples, unassociated with mortality, collected from this bay and from other areas of Tasmania between 1994 and at present. Such samples were generally fixed with 10% formalin in seawater (chosen for practicality for routine use after initial comparison with other fixatives). Selected samples from the initial and 1998 episodes were fixed for transmission electron microscopy (TEM) in 2% glutaraldehyde in seawater or cacodylate buffer, post-fixed in 1% osmium tetroxide and uranyl acetate and dehydrated through alcohol. Samples were then embedded in Spurr’s resin, sectioned at 50–90 nm, stained with uranyl acetate and lead citrate and examined using an Hitachi H300 TEM. Spat too small to remove from shells were decalcified whole for up to 2 h in ‘Fast-Cal’ decalcification solution (Fronine Histo-Labs, NSW, Australia) containing 10% hydrochloric acid and 1% calcium scavenger, until foaming on shaking had almost ceased, before processing between fine mesh foam. Sections were routinely stained with haematoxylin and eosin, though periodic acid-Schiff (PAS), Gram stain, Geimsa and silver stains were also used as required, plus Perls Prussian blue stain to eliminate iron loading as a cause. Standard aquatic animal bacteriology was carried out on representative spat from affected batches.
2.1.3. Mortality 2002: spat collection In May 2002, during an episode of spat mortality at Little Swanport, shellfish were collected from the commonly affected L7–L8 and L1–L2 lease sites and from the nursery site. In addition to histopathological analysis, ten 5–7 mm diameter spat from each site were opened without preservation and shell contents were removed, placed between a microscope slide and coverslip and examined under a Zeiss Axiovert 25 inverted microscope. Mouse bioassays were also conducted on the shell contents of approximately 150 spat (total shell contents = 0.5 g) from each affected site. These spat were opened and shell contents removed to a glass vial and finely chopped using a scalpel blade. An extraction solvent of 2.5 ml of 50% methanol:water:0.2% acetic acid was added and the sample was further homogenised for 2 min. Flesh was removed via centrifugation (1200 g, 10 min) and methanol removed from the supernatant under a stream of nitrogen. The sample was then re-suspended in 2 ml 1% Tween 60. As a control, 150 mm × 5 mm diameter spat were collected from a shellfish nursery 100 km south of the affected site and extracts made in the manner described earlier. Extracts were sent to the Institute of Medical and Veterinary Science (IMVS), Adelaide, for mouse bioassay
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via intraperitoneal injection. Duplicate mice were injected with 1 ml extract each and observed for 24 h. 2.1.4. Algal monitoring An algal monitoring program was implemented in the Little Swanport Estuary following an oyster spat mortality event during 2001. Plankton net (20 m mesh) tows were taken from three lease sites: L7–L8, L1–L2 and NJ (the site from which water is pumped to the land-based shellfish nursery) (Fig. 1). These were taken monthly by staff from the shellfish company between May 2001 and November 2002 and every 1–2 weeks between November 2001 and May 2002. A plankton net was dragged at approximately 1 m below the surface (just above the top of the seagrass beds) concentrating approximately 26 l of water into a 50 ml collecting jar. Plankton samples were preserved in 2% glutaraldehyde and examined under a Zeiss Axiovert 25 microscope for dominant algal species and dinoflagellate cell counts were conducted using Sedgewick–Rafter counting chambers. Percentage spat mortality at each lease was also noted. 2.2. Characterisation of P. rhathymum toxicity 2.2.1. Algal culturing and identification P. rhathymum (isolated from Little Swanport, March 1999) was established in culture prior to use in experimental tests for toxicity. The culture was grown in glass 50 ml flasks in 35 psu GSe medium (Blackburn et al., 1989) at a photon flux density of 60–80 mol photons m−2 s−1 provided by 18 W Sylvania® cool white fluorescent tubes, under a 12:12 h light: dark cycle at 20 ◦ C. Cultures were transferred approximately every 14 days and not considered to be axenic. Cells were examined using an Axioskop 2 Plus light microscope in bright field mode and cells were photographed using a Canon Powershot G1 digital camera. An environmental scanning electron microscope (ESEM) model 2020 was also used for identification, operated at 15 kV in high vacuum mode. For SEM examination, cells were preserved with 1% glutaraldehyde, filtered onto 3 m pore size Nuclepor polycarbonate filter, air dried, mounted on a stub and gold coated.
2.2.2. Artemia salina bioassay Cultured P. rhathymum was assayed for toxicity using a brine shrimp (A. salina) bioassay (Rhodes and Syhre, 1995). Instar II Artemia were used, having been grown from de-capsulated cysts that were inoculated into an aerated tank containing 35 psu seawater at 19 ◦ C under light provided by 18 W Sylvania® cool white fluorescent tubes. Replicates of six Artemia, were placed in culture wells (Iwaki) of 16 mm diameter that were inoculated with 0, 0.01, 0.05, 0.1, 0.25, 0.5, 0.75, 1 and 2 ml P. rhathymum culture (5.91 × 103 cells ml−1 ) and the total volume of each well made up to 2 ml with 35 psu filtered seawater, controls were inoculated with 2 ml GSe culture medium. Using the same methodology, Artemia were also exposed to aliquots of P. rhathymum cell extract. The 2 l culture harvested in late exponential growth phase was centrifuged into two tubes (approximately 6.8 × 106 cells per tube). In one tube an aqueous extraction was obtained, the cells re-suspended in filtered distilled water, ruptured using sonication and liquid nitrogen in three freeze/thaw cycles and then frozen at −80 ◦ C until use. In the second tube methanol was added as a broad spectrum extraction solvent, cells were homogenised and then centrifuged and the supernatant collected. The methanol was removed under a stream of nitrogen and the remnants suspended in 1% Tween 60. Three replicates of six Artemia were exposed to 62.5, 125, 250 and 500 l cell extract in wells as described earlier, and the volume in each well was made up to 2 ml using 35 psu filtered seawater. As a control, Artemia were exposed to the same volumes of 1% Tween 60. 2.2.3. Mouse bioassay A bulk P. rhathymum culture was grown in 1 l GSe media in a 2 l Erlenmeyer flask under conditions indicated previously. The culture was harvested by centrifugation in two tubes, the media supernatant was discarded, leaving approximately 1.8 × 107 cells in each tube. Cell extracts were prepared as described earlier, the water-soluble extract was frozen at −80 ◦ C and the methanol extract was re-suspended in 1% Tween 60. Samples were then transported to the Institute of Medical and Veterinary Science (IMVS), Adelaide. Two female white mice were each injected intraperitoneally with 1 ml extract and observed for 24 h.
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2.2.4. Chattonella marina algal bioassay The toxicity of P. rhathymum against a sensitive microalga, the raphidophyte C. marina was tested (Rhodes, 1994). The experiment was conducted in triplicate 16 mm wells each inoculated with 6.0 × 103 cells of C. marina and volumes of 7.3×103 , 3.7×103 and 7.3 × 102 cells of P. rhathymum. As a control equal volumes of C. marina were inoculated into GSe. Wells were left at a photon flux density of 80 mol photons m−2 s−1 provided by 18 W Sylvania® cool white fluorescent tubes, under a 12:12 h light:dark cycle at 20 ◦ C and each well was examined under a light microscope at 12 and 24 h. After 24 h, if less than 50% total C. marina appeared either dead or morbid, 1 ml from each well was removed and examined. The percentage dead and rounded cells seen in at least 400 C. marina cells were recorded (except in experiment with inoculation of 7.3 × 102 C. marina cells, where at least 100 cells were counted). 2.2.5. Haemolysis assay A haemolysis assay was conducted based on the methods described by Rhodes (1994), which were modified from Edvardsen et al. (1990). Erythrocytes were extracted from farmed Atlantic salmon (Salmo salar) immediately prior to slaughtering for commercial harvest. Half a millilitre of blood was immediately injected into heparin lined VacutainerTM tubes containing 4.5 ml fish RBC diluent. Tubes were maintained at 10 ◦ C until use. Six hundred millilitres of P. rhathymum culture (2.2 × 107 cells) grown in a 1 l glass flask was harvested during late growth phase. The culture was saturated with NaCl and sonicated to aid cell lysis and 300 ml methanol added. Cultures were extracted using dichloromethane following the methods of Rhodes (1994), the final total (300 ml) dichloromethane fraction was dried using a Turbovap II concentration workstation operating at 35 ◦ C, using nitrogen as the evaporation gas. The residue was then re-suspended in 100 l ethanol and 2 ml Tris (to make concentration of 1.5 × 107 cells ml−1 extract, the higher end of the range of 5–15 × 106 cells ml−1 extract described by Edvardsen et al. (1990)). As a control, 600 ml GSe medium was extracted as described earlier. All assays were conducted in 1.5 ml centrifuge tubes. Seven hundred microlitres of P. rhathymum extract and 700 l GSe (control) extract were each added
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to 500 l salmon erythrocytes in fish RBC diluent. As a positive control 700 l Saponin (a commercial haemolysin) was added to 500 l salmon erythrocytes and as a negative control 700 l fish RBC diluent was added to 500 l salmon erythrocytes. These extractions and assays were conducted twice 3 weeks apart using two replicates of 600 ml P. rhathymum culture from Little Swanport. A water-soluble cell extract (2.2 × 106 cells) was also tested. Culture was centrifuged and media supernatant removed, the cells were then re-suspended in 1 ml fish RBC diluent and sonicated for 1 min to lyse cells. Seven hundred microlitres of lysed P. rhathymum cells were then added to 500 l salmon blood. All tubes were refrigerated at 10 ◦ C for 2 h, after which, the condition of the erythrocytes were examined under an Axioskop 2 Plus light microscope in bright field mode. 2.2.6. Oyster spat exposures Experiments were run to investigate the effects of P. rhathymum on the survival of juvenile Pacific oysters (C. gigas). Spat were of similar hatchery origin to those naturally affected but raised in a different nursery location. Spat were approximately 4 mm in shell width and had a wet weight ranging from 0.01 to 0.02 g. P. rhathymum bulk cultures were grown as described previously into two 20 l Nalgene bottles. Cultures were maintained for 2 weeks over the course of the experiment using a semi-continuous culture method, in which 6 l of cells was removed from 18 l of culture every second day and replaced with 6 l fresh GSe media. A growth rate of 0.83 div. per day (established prior to experimentation) allowed for the maintenance of high cell densities. The spat were divided into nine groups of 100 individuals. Each group was placed in an up-welling system made from a section of PVC plumbing pipe (diameter 110 mm, length 100 mm), with one end covered with a 250 m nylon mesh. The spat sat on the mesh and the section of pipe was attached to the side of a 10 l plastic bucket, water was re-circulated within the buckets using aquarium pumps (Aquarium PowerHead 480, Second Nature, NJ, USA) with a flow rate of 700 ml min−1 . Three buckets used as controls were filled with 6 l of 35 ppt unfiltered seawater collected from the Derwent River, Hobart. Three replicate treatment buckets, containing 4 l filtered
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(0.2 m) seawater were inoculated with 2 l of P. rhathymum culture and three additional replicates containing 4 l unfiltered seawater (from the Derwent River, Hobart) were also inoculated with 2 l P. rhathymum culture (as it was thought that the natural mix of food may encourage feeding), final P. rhathymum cell density in each bucket ranged between 5 × 103 and 2.3 × 104 cells ml−1 . This density was comparable to the cell numbers noted in a P. rhathymum bloom (3.1 × 104 cells ml−1 ) by Ismael and Aida (1997) and to the densities of algal cells used in shellfish feeding experiments by Luckenbach et al. (1993). Spat were placed in 0.2 m filtered seawater and not fed for 24 h prior to the start of the experiment. Each day buckets were emptied and replaced with fresh seawater and fresh culture and spat were examined for mortalities (open shells), the mesh was cleaned of any scum every second day. The experiment was run for 21 days and at the conclusion of the experiment, gut
contents of 10 spat were examined under a light microscope to determine whether P. rhathymum had been ingested. The remaining spat were preserved with 10% (v/v) seawater formalin for histopathological examination (methods outlined earlier).
3. Results 3.1. Characterisation of the natural mortality event 3.1.1. Histopathology of oyster spat from natural outbreaks of mortality at Little Swanport Spat examined during each outbreak since 1993 were characterised histologically by overall thinner gut tubule epithelium and the presence of some animals with markedly thinned tubules containing sloughed cells and/or pigment (Fig. 2). Large numbers
Fig. 2. (a) Overview of a section from an oyster spat from Little Swanport undergoing mortality, showing marked thinning and external shell fouling (far right). (b) Higher magnification of (a), showing marked thinning and exfoliation of digestive gland tubule (Dgt) epithelium into lumen (arrow). (c) Sloughed digestive tubules and interstitial tissues where epithelial erosion is complete. (d) Normal (healthy) oyster spat digestive gland tubules, showing star shaped lumen and high absorptive epithelium. All scale bars = 50 m.
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Table 1 Histopathology (percentage of spat affected) recorded from three sites (L7–L8, L1–L2 and NJ) at Little Swanport following a natural mortality event in May 2002 Site
Dead, empty shells
Gill necrosis
Muscle necrosis
Sloughed cells and thinning of digestive tubules
L7–L8 L1–L2 NJ
39 31 Empty shells removed before processing
10 22 –
10 11 –
16 22 19
of bacteria were present in only a few of these sloughed masses, but where gut epithelial erosion was complete this was sometimes seen to lead to deep tissue invasion by bacteria (Fig. 2). No other agents were detected in soft tissues by periodic acid-Schiff, Gram stain, Geimsa stain or acid fast stains, or by electron microscopy. Shells were typically heavily fouled by a number of organisms. No similar gut lesions of marked gut cell sloughing were detected in over 1000 spat examined for routine monitoring from this and other areas at other times during each year. The only other possibly significant finding, irregularly present, was partial hinge ligament erosion associated with fine bacteria morphologically resembling Flavobacterium species. Oysters with this lesion were detected histologically in an early February sample from 1993, but not in samples from January or late February. Similar lesions were an irregular finding in affected batches in 1998, often as only a shallow partial erosion. This finding was more common in May 2000. Oysters with hinge involvement were often moderately to heavily fouled. Occasional shallow erosions were also seen in spat from unaffected areas during routine monitoring. The bacterial flora detected by smears and cultures from the hinge area during various episodes was generally low, mixed, mostly of mixed
Vibrio species, and only occasionally including those of Flavobacterium type. There were negligible bacteria isolated from internal tissues of affected but not dying spat. Changes in other organs were generally considered either too mild to be clearly differentiated from normal variation and sampling damage, or to have occurred in moribund animals with evidence of widespread tissue necrosis. Significant histopathological findings from the examination of spat from each lease site following a severe, natural mortality event in May 2002 are presented in Table 1. As the majority of the oysters were either dead or dying (gill and muscle necrosis is related to moribund oysters), the signs seen were very advanced. 3.1.2. Mortality 2002: gut examination Spat guts examined by direct examination of squashes of fresh spat from affected sites during the May 2002 mortality episode were found to contain predominately P. rhathymum (Fig. 3). At least 20 whole cells P. rhathymum were identified per spat, from the gut of 10 mm × 5 mm spat from sites L1–L2 and L7–L8, and between 10–20 whole cells per spat were identified from 10 mm × 2.5 mm spat from the nursery site. In contrast, P. rhathymum was not seen
Fig. 3. (a) SEM micrograph of P. rhathymum from Little Swanport (scale bar = 5 m). (b) Light micrograph of P. rhathymum from Little Swanport (scale bar = 5 m). (c and d) Evidence of P. rhathymum in gut content of spat from sites L1–L2 and L7–L8, Little Swanport (scale bar = 10 m).
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Fig. 4. Mortality of oyster spat compared with P. rhathymum density at site L7–L8, Little Swanport between May 2001 and May 2002.
upon examination of gut contents from fixed spat, indicating that P. rhathymum was either evacuated or disrupted during spat fixation process. 3.1.3. Mouse bioassay (oyster spat flesh) No symptoms, neurotoxic or hepatotoxic effects were displayed by mice when injected with extracts made using the affected (from the Little Swanport mortality event) or control oyster spat flesh. 3.1.4. Seasonal dinoflagellate distribution in Little Swanport Estuary Spat mortality started to occur at sites L1–L2 and L7–L8 during late January 2002 and continued at levels of 19–88% per tray through to May 2002. Correlation between P. rhathymum density and spat mortality was observed at both sites (Figs. 4 and 5). Other known toxic dinoflagellates O. siamensis, C. monotis, P. lima and Dinophysis species were identified from all sites, however, high numbers never correlated with oyster spat mortality. The only other algal species abundant during this time, was the diatom Chaetoceros (not
shown in Fig. 6), however, this species was seen to bloom at other times of the year without affecting the oysters. An interesting result was the sudden mortality of oyster spat at control site (NJ). Spat deaths had never previously been reported from the nursery until 26 March 2002 when mortality was noted on 1800 and 2240 m sieve size nursery seed. This is also the date at which the phytoplankton net tows suddenly revealed a significant increase in P. rhathymum numbers at the nursery (NJ) site where it had not been recorded previously. 3.2. Characterisation of P. rhathymum toxicity 3.2.1. P. rhathymum taxonomy The cells identified from Little Swanport as P. rhathymum are oval and no pyrenoid is apparent. Cell length is 30–34 m and cell width is 23–27 m (from 25 cells). SEM reveals depressions radiating from the valve centre to form lines perpendicular to the margin and trichocyst pores are located in these. A small spine is located at the anterior end (Fig. 3a).
Fig. 5. Mortality of oyster spat compared with P. rhathymum density at site L1–L2, Little Swanport between May 2001 and May 2002.
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Fig. 6. Dinoflagellate cell density at site L1–L2, Little Swanport, between May 2001 and May 2002.
3.2.2. Artemia bioassay Artemia did not display any adverse effects from exposure to P. rhathymum or P. rhathymum watersoluble extract over 48 h. However, deaths were observed when Artemia was exposed to lipid soluble extracts, in which 60% of Artemia exposed to 250 and 500 l extract died within 24 h, with controls unaffected by identical amounts of Tween 60. No mortality was recorded when exposed to 125 or 62.5 l extract. 3.2.3. Mouse bioassay of P. rhathymum extracts Results of the mouse bioassays are presented in Table 2. Mice died within a short exposure to lipid soluble P. rhathymum extracts. No neurotoxic or hepatotoxic effects were observed.
3.2.4. C. marina bioassay Both P. rhathymum and C. marina appeared to be unaffected by the others presence. The percentage C. marina cells rounding up (very few disintegrated cells were seen) was not significantly different to that seen in the controls. This occurred in all replicates of all densities of C. marina. Average rounded cells (in terms of percentage of C. marina counted) was: control, 8.3 ± 0.0%; C. marina inoculation 7.3 × 103 cells, 10.5 ± 3.5%; C. marina inoculation 3.7 × 103 cells, 12.5 ± 2.2%; C. marina inoculation 7.3 × 102 cells, 9.9 ± 1.58%.
P. rhathymum extract
Result
3.2.5. Haemolysis assay Haemolysis was observed in the salmon erythrocytes exposed to P. rhathymum methanol extract (Fig. 7a–c), however, some haemolysis was also observed in the control extract (GSe media). The haemocytes exposed to Saponin all disintegrated (Fig. 7d), while those exposed to fish RBC diluent remained intact. The water-soluble P. rhathymum extract did not cause haemolysis.
Methanol extract
Mouse (17.7 g): death in 17 min; mouse (19.4 g): death in 19 min Mouse (17.7 g): no death; mouse (19.4 g): death in 26 h
3.2.6. Oyster spat exposure to P. rhathymum Examination of oyster spat gut contents following experimental exposure to P. rhathymum indicated that
Table 2 Mouse response to intraperitoneal injection of aqueous and methanol extracts of P. rhathymum
Aqueous extract
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Fig. 7. Salmon erythrocytes: (a) in fish RBC diluent prior to addition of P. rhathymum methanol extract; (b) immediately after addition of extract; (c) 20 min after addition of extract; (d) erythrocytes when treated with commercial haemolysin Saponin. Scale bar = 10 m. Table 3 Summary of histopathological findings following oyster spat feeding on P. rhathymum (treated) and on unfiltered seawater (control) Symptom
Percentage of treated spata exhibiting symptoms
Percentage of control spatb exhibiting symptoms
Light/occasional
Moderate
High
Total
Light/occasional
Moderate
7.8 13.7 11.8
15.7
2 17.7
3.3 20
3.3
33.3 2
27.5 2
2 15.7 9.8 7.9
23.5 15.7 31.5 15.7 70.6 11.9
43 3.3
Gills Pycnotic cells in gills Phagocytosis through gills Gill reactions
3.7 1.9 5.6
3.7 3.7 20.4
11.2 16.7 20.3
18.6 32.3 46.4
10
Leidig tissue Phagocytosis in Leidig tissue
1.8
18.2
7.3
27.3
Gut Increased haemocytes Presence of brown cells Sloughed cells in gut tubules Phagocytosis in gut region Gut tubules dilated/thin Pycnosis in gut region
a b
3.3
High
Total 3.3 23.3
3.3
46.3 3.3
10 3.3
Percentage of symptoms exhibited in 55 spat exposed to P. rhathymum. Percentage of symptoms exhibited in 30 control spat feeding on unfiltered seawater.
some spat were ingesting considerable cell numbers, but other spat showed minimal ingestion. Dinoflagellate cell number was variable ranging between less than 20 to hundreds of cells per spat in all cases. The predominant symptoms observed in spat fed P. rhathymum were that gut tubules were dilated and thin, many (31.5%) with sloughed cells present in the lumen of the digestive tubules (Fig. 8), additionally, gill reactions, phagocytes and pycnotic cells within the gills were observed (Fig. 9). There was also a high degree of mild gut tubule thinning observed in the control spat (46%), however, this was still lower than the spat exposed to P. rhathymum (70.6%). A summary of significant findings is presented in Table 3 and compared to the controls.
4. Discussion Histological and epidemiological studies have indicated that the oyster spat mortalities seen periodically at Little Swanport occur between January and May, in dry years often following the first significant rainfall. This pattern of mortality and the consistent pathological findings of gut tubule dilation/distension, overall thinner gut tubule epithelium and sloughed epithelium cells have not been observed anywhere else in Tasmania. There is some similarity of these lesions to simple blockage of gut tubules by fragments of crustacea, which evoke a marked host haemocyte response and are seen sporadically in oysters of all age groups. However, serial sectioning of affected spat from batches
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Fig. 8. (a) Digestive gland tubules (Dgt), of spat exposed to a natural mix of phytoplankton, cell epithelium is high and lumens are star shaped indicating a normal, actively feeding and metabolising oyster. (b) Digestive gland tubules of spat exposed to a natural mix of phytoplankton, cell epithelium is thinned possibly indicating mild starvation. (c and d) Examples of the severely dilated digestive gland tubules seen in spat experimentally exposed to P. rhathymum. Arrow indicates an area of epithelial cells sloughed into the lumen of the digestive gland cell. Scale bar = 10 µm.
with mortality did not demonstrate any nidus of tubule blockage. Heavy surface fouling and partial hinge ligament invasion was seen in some batches, but was an inconsistent finding and in most cases the ligament damage was superficial and incomplete. Surface fouling is a normal finding for spat of this size, but it is possible that both the level of fouling and degree of Flavobacterium infection could be enhanced by the environmental factors that are associated with this condition.
Algal monitoring at Little Swanport has established that between late January and May 2002, high density P. rhathymum could be correlated with oyster spat mortality. At this time there was very little else in the water column for the oysters to feed on, except sometimes the diatom Chaetoceros sp. Maximum P. rhathymum density (1.2 × 104 cells l−1 ) was significantly lower than that noted in a P. rhathymum bloom (3.1 × 104 cells ml−1 ) by Ismael and Aida (1997), however, it was the dominant algal species present in
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Fig. 9. Example of oyster gill histopathology when exposed to P. rhathymum, the black arrow indicates an aggregate of haemocytes containing necrotic cells and the white arrows indicate pycnotic cells. Scale bar = 10 µm.
the water column. It is also a semi-epiphytic species and the cell density in the underlying seagrass beds was noted to be higher than that observed from water column samples as collected in this study (unpublished observations). Other known toxin producing dinoflagellates such as Dinophysis spp., P. lima (DSP), C. monotis and O. siamensis were present in the estuary, but never in significant numbers when mortality occurred. Two likely scenarios can be postulated from this data: either P. rhathymum is toxic and is causing the spat mortalities, or P. rhathymum is an indicator species, not affecting the spat themselves, but blooming as a response to the environmental factors that are causing the mortalities. Our experimental studies favour the first hypothesis and we have shown that bloom density P. rhathymum does have a negative impact on oyster spat. Experimental exposure of oyster spat to P. rhathymum resulted in very similar histopathology to that
observed in the Little Swanport casualties. Thinning and dilation of the gut tubules and sloughing of the gut epithelium was noted in many experimental spat. However, as was observed in the natural cases, there was a lack of uniformity of gut morphology and while some spat suffered quite severe effects, others displayed very little significant reaction. The reasons for this are as yet unclear, however, may be due to the natural variation in the initial nutritional condition of the oysters, the number of dinoflagellate cells ingested, or the varying susceptibility of spat to other unidentified stresses. Gut tubule thinning was the most common histopathological finding when oyster spat were exposed to P. rhathymum. This is a symptom often seen as a result of reduced feeding or starvation (Elston, 1999) and mild cases were seen in the control spat, probably due to inadequate phytoplankton densities supplied. As oysters have the ability to close up and stop filtering if toxic stimuli are detected, symptoms resulting
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from starvation, such as gut thinning, may be the first response to high densities of P. rhathymum. If high densities of algae are recognised over a prolonged period of time, the animals may eventually be forced to ingest the cells or starve. Under experimental conditions, when ingestion of P. rhathymum occurred, both gill damage and sloughing of the digestive tubules was seen. Gill damage was not identified as a typical symptom in natural mortality cases, but was common in experimental studies and may be a result of exposure to high densities of P. rhathymum. If the oysters first response to P. rhathymum is to stop feeding, a lag period between the start of a P. rhathymum bloom and oyster symptoms or mortality should be noted in field cases. This was not observed in our algal monitoring results at Little Swanport, however, may have been missed due to sampling frequency or collection technique. Symptoms similar to those seen in this study have been noted by other authors when documenting shellfish responses to toxic algae. Digestive tubule sloughing was noted in scallops fed P. minimum by Wikfors and Smolowitz (1995) and sloughed epithelium cells in the mantle and haemocyte infiltration in the gills were documented by Bricelj et al. (1992) in juvenile oyster (Crassostrea virginica) mortality events correlating to blooms of the dinoflagellate Gymnodinium sanguineum (=Akashiwo sanguinea). Oyster spat mortality was seen in the natural cases but did not occur under experimental conditions with high densities of P. rhathymum. This may be because our experiment was not run for long enough for mortality to be observed or because mortality may be due to the additive effects of stresses experienced in nature. The oysters used in the experimental set-up were obtained from a hatchery where environmental conditions are optimised for fast growth, and thus may have been hardier with more reserve energy supplies. Experiments were also conducted under stable conditions in the laboratory, whereas in leases in the wild, the environment is harsher and shellfish are subject to disease, parasites and abnormal climatic events. Any combination of these may weaken the oysters and hence render them more susceptible to algal toxins. Another possible variable is that cultured P. rhathymum toxicity levels may differ to those in natural populations. Dinoflagellate toxicity can be enhanced or inhibited by particular conditions for example expo-
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sure to specific micronutrients (Yasumoto et al., 1993) or bacterial symbionts (Tosteson et al., 1989). The exact nature of the toxin produced by P. rhathymum is unknown. We have shown that a crude methanolic extract of Tasmanian P. rhathymum results in mouse death in less than 20 min when injected intraperitoneally and this is typical of the fast acting toxins that were isolated from this species by Tindall et al. (1989). However, as no quantification of toxins was reported, direct comparison with the present work is not possible. Tindall et al. (1989) described the toxin as water soluble rather than lipid soluble as identified in our work. In agreement with the current study, work by Nakajima et al. (1981) and Yasumoto et al. (1987) found that the toxins produced by Japanese strains of P. rhathymum were lipid soluble (the water-soluble fraction was not toxic) and contained haemolytic properties. A toxin with such properties was considered to non-specifically attack cell membranes (Nakajima et al., 1981) and this may account for the reactions seen in the affected spat at high densities of algae. However, the haemolytic properties of the Tasmanian strain are questionable, as the raphidophyte C. marina was not affected by exposure to P. rhathymum and a water-soluble cell extract did not cause haemolysis of salmon blood cells. While haemolysis was noted in salmon blood cells exposed to the dichloromethane extract, some haemolysis was also observed in the control (GSe) extract. Thus, results are not conclusive and indicate that there may be something in the media that causes the observed reaction. P. rhathymum is not considered to be a dinoflagellate that would cause poisoning in humans through consumption of toxic shellfish. Mouse bioassay of shellfish flesh was negative, although this was a very small sample of oyster flesh tested and toxin levels may have been too low for detection. Also, blooms/red tides of P. rhathymum have been reported world-wide and no shellfish poisoning incidents have been reported to date. Finally, although mortality was not noted in our experimental set-up, histopathological examination has shown that P. rhathymum does have a detrimental effect on oyster spat. A phytoplankton monitoring program is now being implemented by the shellfish company at Little Swanport to assess P. rhathymum density throughout the year. When P. rhathymum
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numbers increase, stock may be transferred to another area within the estuary or transferred to other outgrow areas in Tasmania.
Acknowledgements Investigation of the Little Swanport mortalities were supported by Project A.2.1 for the Co-operative Research Centre for Aquaculture Limited, and by the Tasmanian Pacific Oyster Health Program. The authors would also like to thank: Greg Hollingsworth and Richard Pugh at Shellfish Culture for assistance with the project and supply of oyster spat, Peter Cameron at IMVS for mouse bioassays and the generosity of Neil Byron at Hobart Pathology and Mick Murfet and Erica Laws at AquaTas. References Blackburn, S., Hallegraeff, G.M., Bolch, C., 1989. Vegetative reproduction and sexual life cycle of the toxic dinoflagellate Gymnodinium catenatum from Tasmania, Australia. J. Phycol. 25, 557–590. Bricelj, V.M., Ford, S.E., Borrero, F.J., Perkins, F.O., Rivara, G., Hillman, R.E., Elston, R.A., Chang, J., 1992. Unexplained mortalities of hatchery-reared, juvenile oysters, Crassostrea virginica (Gmelin). J. Shellfish Res. 11, 331–347. Cortés-Altamirano, R., Sierra-Beltrán, A.P., 2003. Morphology and taxonomy of Prorocentrum mexicanum and reinstatement of Prorocentrum rhathymum (Dinophyceae). J. Phycol. 39, 221– 225. Dickey, R., Faulkner, D., Benesath, F., Andrezehewski, D., 1990. Identification of okadaic acid from a Caribbean dinoflagellate, Prorocentrum concavum. Toxicon 28, 371–378. Department of Primary Industries Water and the Environment (DPIWE), Marine Resources Division, 1998. Marine farming development plan—Great Oyster Bay. Retrieved January 2003 from http://www.dpiwe.tas.gov.au. Edvardsen, B., Moy, F., Paasche, E., 1990. Hemolytic activity in extracts of Chrysochromulina polylepis grown at different levels of selenite and phosphate. In: Graneli, E., Sundstrom, B., Edler, C., Anderson, D.M. (Eds), Toxic Marine Phytoplankton. Elsevier, pp. 284–289. Elston, R.A., 1999. Gastrointestinal epithelial alterations and exfoliation. In: Health Management, Development and Histology of Seed Oysters. World Aquaculture Society, Baton Rouge, LA, Chapter 20, pp. 103–107. Faust, M.A., Larsen, J., Moestrup, Ø., 1999. Potentially toxic phytoplankton. 3. Genus Prorocentrum (Dinophyceae). In: Lindley, J.A. (Ed.), ICES Identification Leaflets for Plankton No. 184. International Council for the Exploration of the Sea, Denmark, pp. 2–23.
Holmes, M., Lewis, R., Jones, A., Wong Hoy, A., 1995. Cooliatoxin, the first toxin from Coolia monotis (Dinophyceae). Nat. Toxins 3, 355–362. Ismael, G.E., Aida, M., 1997. First record of a Prorocentrum rhathymum (Prorocentraceae) red tide in the Gulf of California. Rev. Biol. Trop. 45, 1263. Luckenbach, M.W., Sellner, K.G., Shumway, S.E., Greene, K., 1993. Effects of two bloom forming dinoflagellates Prorocentrum minimum and Gyrodinium uncatenum, on the growth and survival of the eastern oyster, Crassostrea virginica (Gmelin 1791). J. Shellfish Res. 12, 411–415. Morton, S., Bomber, J., Tindall, D., 1994. Environmental effects on the production of okadaic acid from Prorocentrum hoffmannianum Faust. I. Temperature, light and salinity. J. Exp. Mar. Biol. Ecol. 178, 67–77. Nakajima, I., Oshima, Y., Yasumoto, T., 1981. Toxicity of benthic dinoflagellates in Okinawa. Bull. Jpn. Soc. Sci. Fish. 47, 1029– 1033. Pearce, I., Marshall, J.M., Hallegraeff, G.M., 2001. Toxic epiphytic dinoflagellates from east coast Tasmania. In: Hallegraeff, G.M, Blackburn, S.I., Bolch, C.J., Lewis, R.J. (Eds.), Proceedings of the Ninth International Conference on Harmful Algal Blooms. Intergovernmental Oceanographic Commission of UNESCO, pp. 54–57. Rees, C., 1993. Tasmanian seagrass communities. M.Sc. thesis, School of Environmental Studies and Geography, University of Tasmania, pp. 87–90. Rhodes, L.L., 1994. Prymnesiophytes of New Zealand’s coastal waters: taxonomy, physiology and ecology. Ph.D. thesis, Massey University. Rhodes, L.L., Syhre, M., 1995. Okadaic acid production by a New Zealand Prorocentrum lima isolate. N. Z. J. Mar. Freshwater Res. 29, 367–370. Tindall, D., Miller, D., Bomber, J., 1989. Culture and toxicity of dinoflagellates from ciguatera endemic regions of the world. Toxicon 27, 83. Tosteson, T., Ballantine, D., Tosteson, C., Bardales, A., Hensley, V., 1989. Associated bacterial flora, growth and toxicity of cultured benthic dinoflagellates Ostreopsis lenticularis and Gambierdiscus toxicus. Appl. Environ. Microbiol. 41, 137–141. Usami, M., Masayuuki, S., Ishida, S., Inoue, A., Kan, Y., Yasamoto, T., 1995. Palytoxin analogs from the dinoflagellates Ostreopsis siamensis. J. Am. Chem. Soc. 117, 5389–5390. Wikfors, G.H., Smolowitz, R.M., 1995. Experimental and histological studies of four life-history stages of the eastern oyster, Crassostrea virginica, exposed to a cultured strain of the dinoflagellate Prorocentrum minimum. Biol. Bull. 188, 313– 328. Yasumoto, T., Sino, N., Murata, M., 1987. Toxins produced by benthic dinoflagellates. Biol. Bull. 172, 128–131. Yasumoto, T., Satake, M., Fukui, M., Nagai, H., Murata, M., Legrand, A., 1993. A turning point in ciguatera study. In: Smayada, T., Shimizu Y. (Eds.), Toxic Phytoplankton Blooms in the Sea. Elsevier, pp. 455–460. Zhang, Y., Munday, B.L., Handlinger, J.H., 1995. Mass mortality of flat oysters (Ostrea rivularis) associated with a bloom of Prorocentrum sp. in the port of Zhangjiang, South China. Bull. Eur. Assoc. Fish Pathol. 15, 61–63.
Histopathology in Pacific oyster (Crassostrea gigas) spat caused by the dinoflagellate Prorocentrum rhathymum Imojen Pearce a , Judith H. Handlinger b , Gustaaf M. Hallegraeff a,∗ a
b
School of Plant Science, University of Tasmania, Private Bag 55, Hobart 7001, Tasmania, Australia Fish Health Unit, Animal Health Laboratory, Department of Primary Industries Water and the Environment, P.O. Box 46, Kings Meadows 7249, Tasmania, Australia Received 24 August 2003; received in revised form 15 October 2003; accepted 15 November 2003
Abstract The recognition of an apparent association between seasonal oyster spat mortalities (up to 40%) and high Prorocentrum rhathymum density in the Little Swanport Estuary, Tasmania, prompted further experimental investigation into the toxicity by this dinoflagellate. Standard brine shrimp, haemolysis assays and intraperitoneal mouse bioassays revealed fast acting toxins in methanol but not aqueous extracts of P. rhathymum, with mice dying in less than 20 min. Oyster bioassays involved feeding spat (4 mm shell width) for 21 consecutive days on a diet of cultured P. rhathymum at simulated bloom densities (104 cells ml−1 ). No oyster mortality was observed, however, histopathological signs of thin, dilated gut tubules and sloughing of gut cells resembled those seen in affected field samples. In contrast to field samples, gill pathology was also observed in experimental exposure oysters. © 2003 Elsevier B.V. All rights reserved. Keywords: Crassostrea gigas; Histopathology; Oyster; Prorocentrum; Tasmania
1. Introduction A successful hatchery based Pacific oyster (Crassostrea gigas) industry has been established in Tasmania, Australia since the early 1980s. Oyster spat are produced for sale to commercial farmers, initially in a land-based nursery system of flowing unfiltered seawater, and subsequently are transferred to outgrow areas on racks in estuaries. Little Swanport is such a site with leases situated throughout the estuary. The leases are often established over extensive seagrass (Zostera muelleri) beds, the leaf shoots of which support dense ∗ Corresponding author. Tel.: +61-3-62262623; fax: +61-3-62262698. E-mail address: [email protected] (G.M. Hallegraeff).
algal epiphyte growth, particularly in the proximity of the intertidal oyster racks (Rees, 1993). A significant episode of site specific losses of post-hatchery spat was first noted in early January 1993, involving stock from several hatcheries in one of two nursery bays shared by these hatcheries. Smaller similar losses have since occurred at the same site, with notable reoccurrences in May 1998, May–June 2000, May 2001 and March–May 2002. Losses varied between batches present on the site, with maximum losses over several weeks of up to 40% on some racks. Histological, bacteriological and epidemiological study of these outbreaks indicated that the cause was not infectious, and no association could be found with artificial pollutants or pesticides from equipment within the bay or from agricultural or other
1568-9883/$ – see front matter © 2003 Elsevier B.V. All rights reserved. doi:10.1016/j.hal.2003.11.002
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activities upstream (Elston, 1999). However, there was evidence that these losses were more common in low rainfall years, and often followed the first significant rains. The oyster deaths were considered unlikely to be the direct result of freshwater run-off, as spat are usually well able to tolerate reduced salinities, and deaths were not immediate following rains but persisted for up to several weeks, sometimes despite a lack of further rain. The possibility of an origin in soil run-off was also considered. However, while potential areas of acid sulphate soil did exist in the Little Swanport catchment, there was no evidence that these had been drained. Similarly, iron from soil erosion was discounted as no tissue evidence was found in damaged spat. Initial algal studies showed the dinoflagellates Ostreopsis siamensis Schmidt and species of Prorocentrum Ehrenberg were present during the 1998 mortality (Elston, 1999). The involvement of toxic algae was further investigated during 1999, as part of a survey of the potentially toxic epiphytic algal assemblage on the seagrass in the estuary (Pearce et al., 2001). This assemblage included the dinoflagellates O. siamensis, Coolia monotis Meunier, Prorocentrum rhathymum Loeblich, Sherley and Schmidt, and Prorocentrum lima (Ehrenberg) Dodge, strains of which have been shown elsewhere to produce toxins (Holmes et al., 1995; Usami et al., 1995; Yasumoto et al., 1987). Water samples examined during the following periods of high spat mortality (March 2000 and March 2001), revealed that P. rhathymum was blooming on both occasions. This species was initially reported as Prorocentrum mexicanum, but was redefined as P. rhathymum following the redefinition of the two species by Cortés-Altamirano and Sierra-Beltrán (2003). Blooms of P. rhathymum have not previously been reported as causing damage to shellfish. However, the species has been identified as toxic (Tindall et al., 1989; Nakajima et al., 1981) and shellfish mortality has been linked to other members of the genus. Prorocentrum minimum (Pavillard) Schiller has been found to cause death in juvenile oysters and scallops in laboratory feeding experiments (Luckenbach et al., 1993; Wikfors and Smolowitz, 1995), an unidentified Prorocentrum sp. caused mortality of hundreds of flat oysters in China (Zhang et al., 1995) and human poisonings (diarrhetic shellfish poisoning) have been commonly associated with okadaic acid producing
species such as P. lima, Prorocentrum hoffmannianum Faust and Prorocentrum concavum Fukuyo (Dickey et al., 1990; Faust et al., 1999; Morton et al., 1994). In the present work, we aimed to document the characteristics of the recurrent oyster mortality at this site and through an extensive investigation between May 2001 and May 2002, determined whether the Tasmanian strains of P. rhathymum were toxic and responsible for the observed pathology of oyster spat.
2. Methods 2.1. Characterisation of the natural mortality events 2.1.1. Site description Little Swanport is located at the basis of the estuary formed by the Little Swanport River on the western side of Great Oyster Bay, approximately 30 km south of Swansea, Tasmania (Fig. 1). It has a complex hydrography due to its narrow entrance, long channel, numerous shoals and the irregular shape of the estuary. Water circulation is complex, being largely wind driven and is well mixed, as validated by temperature and salinity profiles (DPIWE, 1998). The flushing time is 2.3 tidal cycles or just over a day. Occasionally flooding of the Little Swanport River causes inundation of freshwater into the estuary, however, rainfall rarely exceeds 800 mm per annum (DPIWE, 1998). The surrounding land use is predominantly rural with some residential properties on the northern side of the estuary. 2.1.2. Background: histopathology and health monitoring The health of oysters in the Little Swanport Estuary has been monitored over the past decade through routine health surveillance as part of the Tasmanian Oyster Health Program, and through a Co-operative Research Centre for Aquaculture Project on early mollusc mortality in Tasmania. Histopathology was examined from farm spat undergoing mortality in Little Swanport on archived samples from the early 1993 losses, and samples from mortality episodes in May–June 1998, May 2000 and May 2002. These were compared with routine
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Fig. 1. Study site, Little Swanport, Tasmania (scale bar = 1 km): (1) nursery site (NJ); (2) lease L1–L2; (3) lease L7–L8.
6-monthly surveillance spat samples, unassociated with mortality, collected from this bay and from other areas of Tasmania between 1994 and at present. Such samples were generally fixed with 10% formalin in seawater (chosen for practicality for routine use after initial comparison with other fixatives). Selected samples from the initial and 1998 episodes were fixed for transmission electron microscopy (TEM) in 2% glutaraldehyde in seawater or cacodylate buffer, post-fixed in 1% osmium tetroxide and uranyl acetate and dehydrated through alcohol. Samples were then embedded in Spurr’s resin, sectioned at 50–90 nm, stained with uranyl acetate and lead citrate and examined using an Hitachi H300 TEM. Spat too small to remove from shells were decalcified whole for up to 2 h in ‘Fast-Cal’ decalcification solution (Fronine Histo-Labs, NSW, Australia) containing 10% hydrochloric acid and 1% calcium scavenger, until foaming on shaking had almost ceased, before processing between fine mesh foam. Sections were routinely stained with haematoxylin and eosin, though periodic acid-Schiff (PAS), Gram stain, Geimsa and silver stains were also used as required, plus Perls Prussian blue stain to eliminate iron loading as a cause. Standard aquatic animal bacteriology was carried out on representative spat from affected batches.
2.1.3. Mortality 2002: spat collection In May 2002, during an episode of spat mortality at Little Swanport, shellfish were collected from the commonly affected L7–L8 and L1–L2 lease sites and from the nursery site. In addition to histopathological analysis, ten 5–7 mm diameter spat from each site were opened without preservation and shell contents were removed, placed between a microscope slide and coverslip and examined under a Zeiss Axiovert 25 inverted microscope. Mouse bioassays were also conducted on the shell contents of approximately 150 spat (total shell contents = 0.5 g) from each affected site. These spat were opened and shell contents removed to a glass vial and finely chopped using a scalpel blade. An extraction solvent of 2.5 ml of 50% methanol:water:0.2% acetic acid was added and the sample was further homogenised for 2 min. Flesh was removed via centrifugation (1200 g, 10 min) and methanol removed from the supernatant under a stream of nitrogen. The sample was then re-suspended in 2 ml 1% Tween 60. As a control, 150 mm × 5 mm diameter spat were collected from a shellfish nursery 100 km south of the affected site and extracts made in the manner described earlier. Extracts were sent to the Institute of Medical and Veterinary Science (IMVS), Adelaide, for mouse bioassay
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via intraperitoneal injection. Duplicate mice were injected with 1 ml extract each and observed for 24 h. 2.1.4. Algal monitoring An algal monitoring program was implemented in the Little Swanport Estuary following an oyster spat mortality event during 2001. Plankton net (20 m mesh) tows were taken from three lease sites: L7–L8, L1–L2 and NJ (the site from which water is pumped to the land-based shellfish nursery) (Fig. 1). These were taken monthly by staff from the shellfish company between May 2001 and November 2002 and every 1–2 weeks between November 2001 and May 2002. A plankton net was dragged at approximately 1 m below the surface (just above the top of the seagrass beds) concentrating approximately 26 l of water into a 50 ml collecting jar. Plankton samples were preserved in 2% glutaraldehyde and examined under a Zeiss Axiovert 25 microscope for dominant algal species and dinoflagellate cell counts were conducted using Sedgewick–Rafter counting chambers. Percentage spat mortality at each lease was also noted. 2.2. Characterisation of P. rhathymum toxicity 2.2.1. Algal culturing and identification P. rhathymum (isolated from Little Swanport, March 1999) was established in culture prior to use in experimental tests for toxicity. The culture was grown in glass 50 ml flasks in 35 psu GSe medium (Blackburn et al., 1989) at a photon flux density of 60–80 mol photons m−2 s−1 provided by 18 W Sylvania® cool white fluorescent tubes, under a 12:12 h light: dark cycle at 20 ◦ C. Cultures were transferred approximately every 14 days and not considered to be axenic. Cells were examined using an Axioskop 2 Plus light microscope in bright field mode and cells were photographed using a Canon Powershot G1 digital camera. An environmental scanning electron microscope (ESEM) model 2020 was also used for identification, operated at 15 kV in high vacuum mode. For SEM examination, cells were preserved with 1% glutaraldehyde, filtered onto 3 m pore size Nuclepor polycarbonate filter, air dried, mounted on a stub and gold coated.
2.2.2. Artemia salina bioassay Cultured P. rhathymum was assayed for toxicity using a brine shrimp (A. salina) bioassay (Rhodes and Syhre, 1995). Instar II Artemia were used, having been grown from de-capsulated cysts that were inoculated into an aerated tank containing 35 psu seawater at 19 ◦ C under light provided by 18 W Sylvania® cool white fluorescent tubes. Replicates of six Artemia, were placed in culture wells (Iwaki) of 16 mm diameter that were inoculated with 0, 0.01, 0.05, 0.1, 0.25, 0.5, 0.75, 1 and 2 ml P. rhathymum culture (5.91 × 103 cells ml−1 ) and the total volume of each well made up to 2 ml with 35 psu filtered seawater, controls were inoculated with 2 ml GSe culture medium. Using the same methodology, Artemia were also exposed to aliquots of P. rhathymum cell extract. The 2 l culture harvested in late exponential growth phase was centrifuged into two tubes (approximately 6.8 × 106 cells per tube). In one tube an aqueous extraction was obtained, the cells re-suspended in filtered distilled water, ruptured using sonication and liquid nitrogen in three freeze/thaw cycles and then frozen at −80 ◦ C until use. In the second tube methanol was added as a broad spectrum extraction solvent, cells were homogenised and then centrifuged and the supernatant collected. The methanol was removed under a stream of nitrogen and the remnants suspended in 1% Tween 60. Three replicates of six Artemia were exposed to 62.5, 125, 250 and 500 l cell extract in wells as described earlier, and the volume in each well was made up to 2 ml using 35 psu filtered seawater. As a control, Artemia were exposed to the same volumes of 1% Tween 60. 2.2.3. Mouse bioassay A bulk P. rhathymum culture was grown in 1 l GSe media in a 2 l Erlenmeyer flask under conditions indicated previously. The culture was harvested by centrifugation in two tubes, the media supernatant was discarded, leaving approximately 1.8 × 107 cells in each tube. Cell extracts were prepared as described earlier, the water-soluble extract was frozen at −80 ◦ C and the methanol extract was re-suspended in 1% Tween 60. Samples were then transported to the Institute of Medical and Veterinary Science (IMVS), Adelaide. Two female white mice were each injected intraperitoneally with 1 ml extract and observed for 24 h.
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2.2.4. Chattonella marina algal bioassay The toxicity of P. rhathymum against a sensitive microalga, the raphidophyte C. marina was tested (Rhodes, 1994). The experiment was conducted in triplicate 16 mm wells each inoculated with 6.0 × 103 cells of C. marina and volumes of 7.3×103 , 3.7×103 and 7.3 × 102 cells of P. rhathymum. As a control equal volumes of C. marina were inoculated into GSe. Wells were left at a photon flux density of 80 mol photons m−2 s−1 provided by 18 W Sylvania® cool white fluorescent tubes, under a 12:12 h light:dark cycle at 20 ◦ C and each well was examined under a light microscope at 12 and 24 h. After 24 h, if less than 50% total C. marina appeared either dead or morbid, 1 ml from each well was removed and examined. The percentage dead and rounded cells seen in at least 400 C. marina cells were recorded (except in experiment with inoculation of 7.3 × 102 C. marina cells, where at least 100 cells were counted). 2.2.5. Haemolysis assay A haemolysis assay was conducted based on the methods described by Rhodes (1994), which were modified from Edvardsen et al. (1990). Erythrocytes were extracted from farmed Atlantic salmon (Salmo salar) immediately prior to slaughtering for commercial harvest. Half a millilitre of blood was immediately injected into heparin lined VacutainerTM tubes containing 4.5 ml fish RBC diluent. Tubes were maintained at 10 ◦ C until use. Six hundred millilitres of P. rhathymum culture (2.2 × 107 cells) grown in a 1 l glass flask was harvested during late growth phase. The culture was saturated with NaCl and sonicated to aid cell lysis and 300 ml methanol added. Cultures were extracted using dichloromethane following the methods of Rhodes (1994), the final total (300 ml) dichloromethane fraction was dried using a Turbovap II concentration workstation operating at 35 ◦ C, using nitrogen as the evaporation gas. The residue was then re-suspended in 100 l ethanol and 2 ml Tris (to make concentration of 1.5 × 107 cells ml−1 extract, the higher end of the range of 5–15 × 106 cells ml−1 extract described by Edvardsen et al. (1990)). As a control, 600 ml GSe medium was extracted as described earlier. All assays were conducted in 1.5 ml centrifuge tubes. Seven hundred microlitres of P. rhathymum extract and 700 l GSe (control) extract were each added
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to 500 l salmon erythrocytes in fish RBC diluent. As a positive control 700 l Saponin (a commercial haemolysin) was added to 500 l salmon erythrocytes and as a negative control 700 l fish RBC diluent was added to 500 l salmon erythrocytes. These extractions and assays were conducted twice 3 weeks apart using two replicates of 600 ml P. rhathymum culture from Little Swanport. A water-soluble cell extract (2.2 × 106 cells) was also tested. Culture was centrifuged and media supernatant removed, the cells were then re-suspended in 1 ml fish RBC diluent and sonicated for 1 min to lyse cells. Seven hundred microlitres of lysed P. rhathymum cells were then added to 500 l salmon blood. All tubes were refrigerated at 10 ◦ C for 2 h, after which, the condition of the erythrocytes were examined under an Axioskop 2 Plus light microscope in bright field mode. 2.2.6. Oyster spat exposures Experiments were run to investigate the effects of P. rhathymum on the survival of juvenile Pacific oysters (C. gigas). Spat were of similar hatchery origin to those naturally affected but raised in a different nursery location. Spat were approximately 4 mm in shell width and had a wet weight ranging from 0.01 to 0.02 g. P. rhathymum bulk cultures were grown as described previously into two 20 l Nalgene bottles. Cultures were maintained for 2 weeks over the course of the experiment using a semi-continuous culture method, in which 6 l of cells was removed from 18 l of culture every second day and replaced with 6 l fresh GSe media. A growth rate of 0.83 div. per day (established prior to experimentation) allowed for the maintenance of high cell densities. The spat were divided into nine groups of 100 individuals. Each group was placed in an up-welling system made from a section of PVC plumbing pipe (diameter 110 mm, length 100 mm), with one end covered with a 250 m nylon mesh. The spat sat on the mesh and the section of pipe was attached to the side of a 10 l plastic bucket, water was re-circulated within the buckets using aquarium pumps (Aquarium PowerHead 480, Second Nature, NJ, USA) with a flow rate of 700 ml min−1 . Three buckets used as controls were filled with 6 l of 35 ppt unfiltered seawater collected from the Derwent River, Hobart. Three replicate treatment buckets, containing 4 l filtered
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(0.2 m) seawater were inoculated with 2 l of P. rhathymum culture and three additional replicates containing 4 l unfiltered seawater (from the Derwent River, Hobart) were also inoculated with 2 l P. rhathymum culture (as it was thought that the natural mix of food may encourage feeding), final P. rhathymum cell density in each bucket ranged between 5 × 103 and 2.3 × 104 cells ml−1 . This density was comparable to the cell numbers noted in a P. rhathymum bloom (3.1 × 104 cells ml−1 ) by Ismael and Aida (1997) and to the densities of algal cells used in shellfish feeding experiments by Luckenbach et al. (1993). Spat were placed in 0.2 m filtered seawater and not fed for 24 h prior to the start of the experiment. Each day buckets were emptied and replaced with fresh seawater and fresh culture and spat were examined for mortalities (open shells), the mesh was cleaned of any scum every second day. The experiment was run for 21 days and at the conclusion of the experiment, gut
contents of 10 spat were examined under a light microscope to determine whether P. rhathymum had been ingested. The remaining spat were preserved with 10% (v/v) seawater formalin for histopathological examination (methods outlined earlier).
3. Results 3.1. Characterisation of the natural mortality event 3.1.1. Histopathology of oyster spat from natural outbreaks of mortality at Little Swanport Spat examined during each outbreak since 1993 were characterised histologically by overall thinner gut tubule epithelium and the presence of some animals with markedly thinned tubules containing sloughed cells and/or pigment (Fig. 2). Large numbers
Fig. 2. (a) Overview of a section from an oyster spat from Little Swanport undergoing mortality, showing marked thinning and external shell fouling (far right). (b) Higher magnification of (a), showing marked thinning and exfoliation of digestive gland tubule (Dgt) epithelium into lumen (arrow). (c) Sloughed digestive tubules and interstitial tissues where epithelial erosion is complete. (d) Normal (healthy) oyster spat digestive gland tubules, showing star shaped lumen and high absorptive epithelium. All scale bars = 50 m.
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Table 1 Histopathology (percentage of spat affected) recorded from three sites (L7–L8, L1–L2 and NJ) at Little Swanport following a natural mortality event in May 2002 Site
Dead, empty shells
Gill necrosis
Muscle necrosis
Sloughed cells and thinning of digestive tubules
L7–L8 L1–L2 NJ
39 31 Empty shells removed before processing
10 22 –
10 11 –
16 22 19
of bacteria were present in only a few of these sloughed masses, but where gut epithelial erosion was complete this was sometimes seen to lead to deep tissue invasion by bacteria (Fig. 2). No other agents were detected in soft tissues by periodic acid-Schiff, Gram stain, Geimsa stain or acid fast stains, or by electron microscopy. Shells were typically heavily fouled by a number of organisms. No similar gut lesions of marked gut cell sloughing were detected in over 1000 spat examined for routine monitoring from this and other areas at other times during each year. The only other possibly significant finding, irregularly present, was partial hinge ligament erosion associated with fine bacteria morphologically resembling Flavobacterium species. Oysters with this lesion were detected histologically in an early February sample from 1993, but not in samples from January or late February. Similar lesions were an irregular finding in affected batches in 1998, often as only a shallow partial erosion. This finding was more common in May 2000. Oysters with hinge involvement were often moderately to heavily fouled. Occasional shallow erosions were also seen in spat from unaffected areas during routine monitoring. The bacterial flora detected by smears and cultures from the hinge area during various episodes was generally low, mixed, mostly of mixed
Vibrio species, and only occasionally including those of Flavobacterium type. There were negligible bacteria isolated from internal tissues of affected but not dying spat. Changes in other organs were generally considered either too mild to be clearly differentiated from normal variation and sampling damage, or to have occurred in moribund animals with evidence of widespread tissue necrosis. Significant histopathological findings from the examination of spat from each lease site following a severe, natural mortality event in May 2002 are presented in Table 1. As the majority of the oysters were either dead or dying (gill and muscle necrosis is related to moribund oysters), the signs seen were very advanced. 3.1.2. Mortality 2002: gut examination Spat guts examined by direct examination of squashes of fresh spat from affected sites during the May 2002 mortality episode were found to contain predominately P. rhathymum (Fig. 3). At least 20 whole cells P. rhathymum were identified per spat, from the gut of 10 mm × 5 mm spat from sites L1–L2 and L7–L8, and between 10–20 whole cells per spat were identified from 10 mm × 2.5 mm spat from the nursery site. In contrast, P. rhathymum was not seen
Fig. 3. (a) SEM micrograph of P. rhathymum from Little Swanport (scale bar = 5 m). (b) Light micrograph of P. rhathymum from Little Swanport (scale bar = 5 m). (c and d) Evidence of P. rhathymum in gut content of spat from sites L1–L2 and L7–L8, Little Swanport (scale bar = 10 m).
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Fig. 4. Mortality of oyster spat compared with P. rhathymum density at site L7–L8, Little Swanport between May 2001 and May 2002.
upon examination of gut contents from fixed spat, indicating that P. rhathymum was either evacuated or disrupted during spat fixation process. 3.1.3. Mouse bioassay (oyster spat flesh) No symptoms, neurotoxic or hepatotoxic effects were displayed by mice when injected with extracts made using the affected (from the Little Swanport mortality event) or control oyster spat flesh. 3.1.4. Seasonal dinoflagellate distribution in Little Swanport Estuary Spat mortality started to occur at sites L1–L2 and L7–L8 during late January 2002 and continued at levels of 19–88% per tray through to May 2002. Correlation between P. rhathymum density and spat mortality was observed at both sites (Figs. 4 and 5). Other known toxic dinoflagellates O. siamensis, C. monotis, P. lima and Dinophysis species were identified from all sites, however, high numbers never correlated with oyster spat mortality. The only other algal species abundant during this time, was the diatom Chaetoceros (not
shown in Fig. 6), however, this species was seen to bloom at other times of the year without affecting the oysters. An interesting result was the sudden mortality of oyster spat at control site (NJ). Spat deaths had never previously been reported from the nursery until 26 March 2002 when mortality was noted on 1800 and 2240 m sieve size nursery seed. This is also the date at which the phytoplankton net tows suddenly revealed a significant increase in P. rhathymum numbers at the nursery (NJ) site where it had not been recorded previously. 3.2. Characterisation of P. rhathymum toxicity 3.2.1. P. rhathymum taxonomy The cells identified from Little Swanport as P. rhathymum are oval and no pyrenoid is apparent. Cell length is 30–34 m and cell width is 23–27 m (from 25 cells). SEM reveals depressions radiating from the valve centre to form lines perpendicular to the margin and trichocyst pores are located in these. A small spine is located at the anterior end (Fig. 3a).
Fig. 5. Mortality of oyster spat compared with P. rhathymum density at site L1–L2, Little Swanport between May 2001 and May 2002.
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Fig. 6. Dinoflagellate cell density at site L1–L2, Little Swanport, between May 2001 and May 2002.
3.2.2. Artemia bioassay Artemia did not display any adverse effects from exposure to P. rhathymum or P. rhathymum watersoluble extract over 48 h. However, deaths were observed when Artemia was exposed to lipid soluble extracts, in which 60% of Artemia exposed to 250 and 500 l extract died within 24 h, with controls unaffected by identical amounts of Tween 60. No mortality was recorded when exposed to 125 or 62.5 l extract. 3.2.3. Mouse bioassay of P. rhathymum extracts Results of the mouse bioassays are presented in Table 2. Mice died within a short exposure to lipid soluble P. rhathymum extracts. No neurotoxic or hepatotoxic effects were observed.
3.2.4. C. marina bioassay Both P. rhathymum and C. marina appeared to be unaffected by the others presence. The percentage C. marina cells rounding up (very few disintegrated cells were seen) was not significantly different to that seen in the controls. This occurred in all replicates of all densities of C. marina. Average rounded cells (in terms of percentage of C. marina counted) was: control, 8.3 ± 0.0%; C. marina inoculation 7.3 × 103 cells, 10.5 ± 3.5%; C. marina inoculation 3.7 × 103 cells, 12.5 ± 2.2%; C. marina inoculation 7.3 × 102 cells, 9.9 ± 1.58%.
P. rhathymum extract
Result
3.2.5. Haemolysis assay Haemolysis was observed in the salmon erythrocytes exposed to P. rhathymum methanol extract (Fig. 7a–c), however, some haemolysis was also observed in the control extract (GSe media). The haemocytes exposed to Saponin all disintegrated (Fig. 7d), while those exposed to fish RBC diluent remained intact. The water-soluble P. rhathymum extract did not cause haemolysis.
Methanol extract
Mouse (17.7 g): death in 17 min; mouse (19.4 g): death in 19 min Mouse (17.7 g): no death; mouse (19.4 g): death in 26 h
3.2.6. Oyster spat exposure to P. rhathymum Examination of oyster spat gut contents following experimental exposure to P. rhathymum indicated that
Table 2 Mouse response to intraperitoneal injection of aqueous and methanol extracts of P. rhathymum
Aqueous extract
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Fig. 7. Salmon erythrocytes: (a) in fish RBC diluent prior to addition of P. rhathymum methanol extract; (b) immediately after addition of extract; (c) 20 min after addition of extract; (d) erythrocytes when treated with commercial haemolysin Saponin. Scale bar = 10 m. Table 3 Summary of histopathological findings following oyster spat feeding on P. rhathymum (treated) and on unfiltered seawater (control) Symptom
Percentage of treated spata exhibiting symptoms
Percentage of control spatb exhibiting symptoms
Light/occasional
Moderate
High
Total
Light/occasional
Moderate
7.8 13.7 11.8
15.7
2 17.7
3.3 20
3.3
33.3 2
27.5 2
2 15.7 9.8 7.9
23.5 15.7 31.5 15.7 70.6 11.9
43 3.3
Gills Pycnotic cells in gills Phagocytosis through gills Gill reactions
3.7 1.9 5.6
3.7 3.7 20.4
11.2 16.7 20.3
18.6 32.3 46.4
10
Leidig tissue Phagocytosis in Leidig tissue
1.8
18.2
7.3
27.3
Gut Increased haemocytes Presence of brown cells Sloughed cells in gut tubules Phagocytosis in gut region Gut tubules dilated/thin Pycnosis in gut region
a b
3.3
High
Total 3.3 23.3
3.3
46.3 3.3
10 3.3
Percentage of symptoms exhibited in 55 spat exposed to P. rhathymum. Percentage of symptoms exhibited in 30 control spat feeding on unfiltered seawater.
some spat were ingesting considerable cell numbers, but other spat showed minimal ingestion. Dinoflagellate cell number was variable ranging between less than 20 to hundreds of cells per spat in all cases. The predominant symptoms observed in spat fed P. rhathymum were that gut tubules were dilated and thin, many (31.5%) with sloughed cells present in the lumen of the digestive tubules (Fig. 8), additionally, gill reactions, phagocytes and pycnotic cells within the gills were observed (Fig. 9). There was also a high degree of mild gut tubule thinning observed in the control spat (46%), however, this was still lower than the spat exposed to P. rhathymum (70.6%). A summary of significant findings is presented in Table 3 and compared to the controls.
4. Discussion Histological and epidemiological studies have indicated that the oyster spat mortalities seen periodically at Little Swanport occur between January and May, in dry years often following the first significant rainfall. This pattern of mortality and the consistent pathological findings of gut tubule dilation/distension, overall thinner gut tubule epithelium and sloughed epithelium cells have not been observed anywhere else in Tasmania. There is some similarity of these lesions to simple blockage of gut tubules by fragments of crustacea, which evoke a marked host haemocyte response and are seen sporadically in oysters of all age groups. However, serial sectioning of affected spat from batches
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Fig. 8. (a) Digestive gland tubules (Dgt), of spat exposed to a natural mix of phytoplankton, cell epithelium is high and lumens are star shaped indicating a normal, actively feeding and metabolising oyster. (b) Digestive gland tubules of spat exposed to a natural mix of phytoplankton, cell epithelium is thinned possibly indicating mild starvation. (c and d) Examples of the severely dilated digestive gland tubules seen in spat experimentally exposed to P. rhathymum. Arrow indicates an area of epithelial cells sloughed into the lumen of the digestive gland cell. Scale bar = 10 µm.
with mortality did not demonstrate any nidus of tubule blockage. Heavy surface fouling and partial hinge ligament invasion was seen in some batches, but was an inconsistent finding and in most cases the ligament damage was superficial and incomplete. Surface fouling is a normal finding for spat of this size, but it is possible that both the level of fouling and degree of Flavobacterium infection could be enhanced by the environmental factors that are associated with this condition.
Algal monitoring at Little Swanport has established that between late January and May 2002, high density P. rhathymum could be correlated with oyster spat mortality. At this time there was very little else in the water column for the oysters to feed on, except sometimes the diatom Chaetoceros sp. Maximum P. rhathymum density (1.2 × 104 cells l−1 ) was significantly lower than that noted in a P. rhathymum bloom (3.1 × 104 cells ml−1 ) by Ismael and Aida (1997), however, it was the dominant algal species present in
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Fig. 9. Example of oyster gill histopathology when exposed to P. rhathymum, the black arrow indicates an aggregate of haemocytes containing necrotic cells and the white arrows indicate pycnotic cells. Scale bar = 10 µm.
the water column. It is also a semi-epiphytic species and the cell density in the underlying seagrass beds was noted to be higher than that observed from water column samples as collected in this study (unpublished observations). Other known toxin producing dinoflagellates such as Dinophysis spp., P. lima (DSP), C. monotis and O. siamensis were present in the estuary, but never in significant numbers when mortality occurred. Two likely scenarios can be postulated from this data: either P. rhathymum is toxic and is causing the spat mortalities, or P. rhathymum is an indicator species, not affecting the spat themselves, but blooming as a response to the environmental factors that are causing the mortalities. Our experimental studies favour the first hypothesis and we have shown that bloom density P. rhathymum does have a negative impact on oyster spat. Experimental exposure of oyster spat to P. rhathymum resulted in very similar histopathology to that
observed in the Little Swanport casualties. Thinning and dilation of the gut tubules and sloughing of the gut epithelium was noted in many experimental spat. However, as was observed in the natural cases, there was a lack of uniformity of gut morphology and while some spat suffered quite severe effects, others displayed very little significant reaction. The reasons for this are as yet unclear, however, may be due to the natural variation in the initial nutritional condition of the oysters, the number of dinoflagellate cells ingested, or the varying susceptibility of spat to other unidentified stresses. Gut tubule thinning was the most common histopathological finding when oyster spat were exposed to P. rhathymum. This is a symptom often seen as a result of reduced feeding or starvation (Elston, 1999) and mild cases were seen in the control spat, probably due to inadequate phytoplankton densities supplied. As oysters have the ability to close up and stop filtering if toxic stimuli are detected, symptoms resulting
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from starvation, such as gut thinning, may be the first response to high densities of P. rhathymum. If high densities of algae are recognised over a prolonged period of time, the animals may eventually be forced to ingest the cells or starve. Under experimental conditions, when ingestion of P. rhathymum occurred, both gill damage and sloughing of the digestive tubules was seen. Gill damage was not identified as a typical symptom in natural mortality cases, but was common in experimental studies and may be a result of exposure to high densities of P. rhathymum. If the oysters first response to P. rhathymum is to stop feeding, a lag period between the start of a P. rhathymum bloom and oyster symptoms or mortality should be noted in field cases. This was not observed in our algal monitoring results at Little Swanport, however, may have been missed due to sampling frequency or collection technique. Symptoms similar to those seen in this study have been noted by other authors when documenting shellfish responses to toxic algae. Digestive tubule sloughing was noted in scallops fed P. minimum by Wikfors and Smolowitz (1995) and sloughed epithelium cells in the mantle and haemocyte infiltration in the gills were documented by Bricelj et al. (1992) in juvenile oyster (Crassostrea virginica) mortality events correlating to blooms of the dinoflagellate Gymnodinium sanguineum (=Akashiwo sanguinea). Oyster spat mortality was seen in the natural cases but did not occur under experimental conditions with high densities of P. rhathymum. This may be because our experiment was not run for long enough for mortality to be observed or because mortality may be due to the additive effects of stresses experienced in nature. The oysters used in the experimental set-up were obtained from a hatchery where environmental conditions are optimised for fast growth, and thus may have been hardier with more reserve energy supplies. Experiments were also conducted under stable conditions in the laboratory, whereas in leases in the wild, the environment is harsher and shellfish are subject to disease, parasites and abnormal climatic events. Any combination of these may weaken the oysters and hence render them more susceptible to algal toxins. Another possible variable is that cultured P. rhathymum toxicity levels may differ to those in natural populations. Dinoflagellate toxicity can be enhanced or inhibited by particular conditions for example expo-
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sure to specific micronutrients (Yasumoto et al., 1993) or bacterial symbionts (Tosteson et al., 1989). The exact nature of the toxin produced by P. rhathymum is unknown. We have shown that a crude methanolic extract of Tasmanian P. rhathymum results in mouse death in less than 20 min when injected intraperitoneally and this is typical of the fast acting toxins that were isolated from this species by Tindall et al. (1989). However, as no quantification of toxins was reported, direct comparison with the present work is not possible. Tindall et al. (1989) described the toxin as water soluble rather than lipid soluble as identified in our work. In agreement with the current study, work by Nakajima et al. (1981) and Yasumoto et al. (1987) found that the toxins produced by Japanese strains of P. rhathymum were lipid soluble (the water-soluble fraction was not toxic) and contained haemolytic properties. A toxin with such properties was considered to non-specifically attack cell membranes (Nakajima et al., 1981) and this may account for the reactions seen in the affected spat at high densities of algae. However, the haemolytic properties of the Tasmanian strain are questionable, as the raphidophyte C. marina was not affected by exposure to P. rhathymum and a water-soluble cell extract did not cause haemolysis of salmon blood cells. While haemolysis was noted in salmon blood cells exposed to the dichloromethane extract, some haemolysis was also observed in the control (GSe) extract. Thus, results are not conclusive and indicate that there may be something in the media that causes the observed reaction. P. rhathymum is not considered to be a dinoflagellate that would cause poisoning in humans through consumption of toxic shellfish. Mouse bioassay of shellfish flesh was negative, although this was a very small sample of oyster flesh tested and toxin levels may have been too low for detection. Also, blooms/red tides of P. rhathymum have been reported world-wide and no shellfish poisoning incidents have been reported to date. Finally, although mortality was not noted in our experimental set-up, histopathological examination has shown that P. rhathymum does have a detrimental effect on oyster spat. A phytoplankton monitoring program is now being implemented by the shellfish company at Little Swanport to assess P. rhathymum density throughout the year. When P. rhathymum
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numbers increase, stock may be transferred to another area within the estuary or transferred to other outgrow areas in Tasmania.
Acknowledgements Investigation of the Little Swanport mortalities were supported by Project A.2.1 for the Co-operative Research Centre for Aquaculture Limited, and by the Tasmanian Pacific Oyster Health Program. The authors would also like to thank: Greg Hollingsworth and Richard Pugh at Shellfish Culture for assistance with the project and supply of oyster spat, Peter Cameron at IMVS for mouse bioassays and the generosity of Neil Byron at Hobart Pathology and Mick Murfet and Erica Laws at AquaTas. References Blackburn, S., Hallegraeff, G.M., Bolch, C., 1989. Vegetative reproduction and sexual life cycle of the toxic dinoflagellate Gymnodinium catenatum from Tasmania, Australia. J. Phycol. 25, 557–590. Bricelj, V.M., Ford, S.E., Borrero, F.J., Perkins, F.O., Rivara, G., Hillman, R.E., Elston, R.A., Chang, J., 1992. Unexplained mortalities of hatchery-reared, juvenile oysters, Crassostrea virginica (Gmelin). J. Shellfish Res. 11, 331–347. Cortés-Altamirano, R., Sierra-Beltrán, A.P., 2003. Morphology and taxonomy of Prorocentrum mexicanum and reinstatement of Prorocentrum rhathymum (Dinophyceae). J. Phycol. 39, 221– 225. Dickey, R., Faulkner, D., Benesath, F., Andrezehewski, D., 1990. Identification of okadaic acid from a Caribbean dinoflagellate, Prorocentrum concavum. Toxicon 28, 371–378. Department of Primary Industries Water and the Environment (DPIWE), Marine Resources Division, 1998. Marine farming development plan—Great Oyster Bay. Retrieved January 2003 from http://www.dpiwe.tas.gov.au. Edvardsen, B., Moy, F., Paasche, E., 1990. Hemolytic activity in extracts of Chrysochromulina polylepis grown at different levels of selenite and phosphate. In: Graneli, E., Sundstrom, B., Edler, C., Anderson, D.M. (Eds), Toxic Marine Phytoplankton. Elsevier, pp. 284–289. Elston, R.A., 1999. Gastrointestinal epithelial alterations and exfoliation. In: Health Management, Development and Histology of Seed Oysters. World Aquaculture Society, Baton Rouge, LA, Chapter 20, pp. 103–107. Faust, M.A., Larsen, J., Moestrup, Ø., 1999. Potentially toxic phytoplankton. 3. Genus Prorocentrum (Dinophyceae). In: Lindley, J.A. (Ed.), ICES Identification Leaflets for Plankton No. 184. International Council for the Exploration of the Sea, Denmark, pp. 2–23.
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