Printer Friendly

Differential retention of okadaic acid (OA) group toxins and pectenotoxins (PTX) in the blue mussel, Mytilus edulis (L.), and European flat oyster, Ostrea edulis (L.).

ABSTRACT This paper presents the results from an experiment, where uptake and elimination of diarrhetic shellfish toxins (DST) of the okadaic acid (OA) and pectenotoxin (PTX) groups were compared between blue mussels (Mytilus edulis) and European oysters (Ostrea edulis). Caged mussels and oysters were suspended in the water column and exposed to a dense bloom of Dinophysis acuta (500-2000 cells [L.sup.-1]) for 4 wk, which was followed by detoxification in the laboratory during 7 wk. Weekly sampling and analysis of OA-group toxins including fatty acid esters ('DTX-3') as well as PTX in individual shellfish and plankton samples were performed. The results showed that mussels rapidly accumulated OA-group toxins to levels about 10 times above the regulation limit (160 [micro]g OA kg-1 mussel) whereas concentrations never reached this limit in oysters during the field exposure. Overall, levels were 10-50 times greater in mussels. The OA-group toxins were mainly in the form of esters (>90%) in oysters, whereas in mussels, the esters constituted only a minor proportion of total OA toxin levels. Reduction rates were estimated for each OA toxin to evaluate if faster elimination could explain the lower toxin retention in oysters. However, no consistent species-specific difference in reduction rates were observed, but esters of OA appeared to be reduced at a faster rate in oysters ([t.sub.1/2] = 23 days) compared with mussels ([t.sub.1/2] = 35 days). In both species, the free form of OA was eliminated at a faster rate ([t.sub.1/2] = 15-17 days) compared with free DTX1 ([t.sub.1/2] = 23-31 days) and DTX2 ([t.sub.1/2] = 28-33 days). Slightly slower elimination rates were estimated for the ester forms ([t.sub.1/2] = 23-42 days). Regarding PTX, PTX2 seco acid (PTX2 SA) was the major PTX detected in both species, but small amounts of PTX2, PTX 12 and PTX 12 SA were also found. As for the OA-group toxins, oysters generally contained lower total amounts of PTX compared with mussels, but the difference was much less apparent. Estimation of reduction rates of the different PTX compounds showed that these toxins were rapidly eliminated in both oysters and mussels ([t.sub.1/2] = 6- 13 days). We propose that differential rates of gut assimilation and/or biotransformation of the OA-group and PTX explain some of the observed differences in retention and toxin profiles between the bivalves, rather than differences in elimination rates. However, models related to differences in feeding rates, particle selection and behavioural response to toxic algae should be tested in future experiments to evaluate the importance of preingestive mechanisms to the differential toxin retention in these bivalves. From the industrial perspective, our results suggest that O. edulis may be regarded as a low-risk species for DST contamination, which should be taken into consideration by regulatory authorities in charge of sampling frequencies and monitoring programs for shellfish toxins.

KEY WORDS: Mytilus edulis, Ostrea edulis, Skagerrack, diarrhetic shellfish toxins, DSP, okadaic acid, pectenotoxins, elimination


Diarrhetic shellfish poisoning (DSP) is a globally distributed seafood poisoning connected to consumption of shellfish contaminated by algal toxins, which originate from dinoflagellates of the genera Dinophysis and Prorocentrum (Yasumoto et al. 1978, Daranas et al. 2001, Van Dolah 2000). In Sweden and Norway, DSP toxins (DST) of the okadaic acid (OA) group were first discovered in 1983 in blue mussels, Mytilus edulis L. (Krogh et al. 1985). Since then, it has been shown that mussels recurrently accumulate levels of OA-toxins above the EU regulation limit (160 [micro]g OA equivalents [kg.sup.-1] shellfish meat) during autumn and winter months, causing prolonged periods of harvest prohibitions (Karlson et al. 2007). This problem has to some extent reduced the possibilities for a major expansion of the mussel farming industry in the Nordic countries, which otherwise have excellent coastal and hydrographical conditions for mussel aquaculture.

Fisheries for the native European oyster, Ostrea edulis (L.), occur along the Swedish and Norwegian coastline and also in Danish waters. This is a seafood product of high commercial value, much because of the fact that these populations are free from the deleterious parasite infections (Bonamiose and Marteilliose), which have caused devastating effects on O. edulis in other parts of Europe. The natural population sizes are, however, small but recent introduction of oysters hatcheries are expected to increase the production of O. edulis in Scandinavia.

Oysters are regularly monitored for biotoxins, but because it is still a relatively small industry in these countries, few samples are analyzed yearly. Nevertheless, monitoring of lipophilic shellfish toxins in Sweden has shown that oysters rarely contain OA-toxins above the detection limit of the chemical HPLC-MS analysis (>10 [micro]g [kg.sup.-1] shellfish meat) (Swedish National Food Administration, shellfish control program, Difference in exposure to the toxic algae is a likely explanation for this observed pattern, because mussels from long-line farms and native oysters do not occupy the same habitats. However, results from a field study performed in 1999, where co-occurring natural populations of mussels and oysters were sampled at several places along the Swedish west coast, revealed that mussels contained OA around 160 [micro]g [kg.sup.-1] shellfish, whereas OA in all oysters was below the detection limit (Svensson et al. 2000). This was somewhat puzzling results, which suggested that oysters do not retain OA-toxins. The biotransformation products, (i.e., fatty acid esters of OA, often referred to as "DTX-3") were, however, not analyzed and the possibility remains that OA occurred mainly in esterified forms in the oysters.

The objective of the present study has been to follow up the results from the 1999 study to test the generality of the observation regarding the interspecies differences in accumulation of DST. An experiment was conducted during the autumn of 2005, where mussels and oysters were collected, placed in cages and exposed to toxic algae for 4 wk during a bloom event of Dinophysis spp., producing DSP toxins of both the OA and pectenotoxin (PTX) group (Yasumoto et al. 1989, Lindahl et al. 2007). This was followed by a detoxification period in controlled laboratory conditions for 7 wk. Uptake and detoxification of the various toxin components within the OA and PTX group of toxins were studied. Based on our previous observations, it was predicted that (1). in general, the total toxin content would be lower in oysters compared with mussels and (2). if detected, the ester forms of the OA-group toxins would predominate in the oysters. This paper presents the first records of PTX in Swedish shellfish. Also, to our knowledge, this is the first time a comparison between the dynamics of the parent toxins of both groups produced by Dinophysis spp. have been done in M. edulis and O. edulis


Experimental Setup and Sampling

Field Exposure

The field exposure stage of the experiment was carried out between September 3 and October l, 2005 in Skagerrack, in a fjord northwest of the island of Orust on the Swedish west coast. The location (=Bjornsund, N 58[degrees]12'5"; E 11[degrees]30'3") is a 9 m. deep sill sound between the Koljo fjord and coastal waters and this area is regularly subjected to blooms of Dinophysis acuta (Ehrenberg) during the autumn months. Blue mussels from a local long-line farm in Northern Bohuslan (shell length 7-8 cm, soft tissue wet weight 10.0 g [+ or -] 3.3 g standard deviation SD) and native oysters collected at 3-6 m depth in the vicinity of Tjarno Marine Biological Laboratory (shell length 8-9 cm, soft tissue wet weight 11.6 g [+ or -] 2.7 g SD) were collected by diving a few days prior to the start. The bivalves were taken to Tjarno Laboratory where they were cleaned and placed in plastic cages (40(1) * 30(w) * 10(h) cm) with a density of 20 mussels and 10 oysters respectively in each cage. A lower density of the oysters was chosen to avoid contact between the oysters. A total of 60 individuals of each species were put out at the start of the experiment. The cages were hung in the water column at 3 m depth. Mussels and oysters were sampled weekly during the field exposure period. At each sampling, 5 individuals of mussels and oysters respectively were removed from the cages and transported to the laboratory where they were chucked and frozen for subsequent toxin analysis. Weekly collection of water samples for analysis of presence of Dinophysis spp. and determination of toxin levels was also done at each sampling occasion. One-hundred liters were pumped from 3 m through a 20 gm plankton net. The concentrate was transferred to 50 mL test tubes and 1 mL was removed and fixed in 1% Lugol solutions for algal identification and enumeration. The remaining plankton solution was filtered onto a GF/F glass fiber filter in the laboratory and frozen until analysis of DST was done.

Laboratory Detoxification

On October 1, the remaining mussels and oysters were transported to Tjarno laboratory and placed in plastic containers in a flow through system with filtered (0.45 [micro]m) seawater from 37 m depth. Additional food was not supplied during detoxification. The shellfish were kept in this system until November 18 when the experiment was terminated. Five mussels and oysters were sampled weekly for toxin analysis during this period.

Sample Preparation for Analysis of DST in Shellfish and Plankton

For toxin analysis, mussels and oysters were thawed and drained and the soft flesh was then removed from the shell. The hepatopancreas (HP) was separated from the rest of the flesh and the weight of both parts was recorded for calculation of % HP. Toxins were analyzed in the HP tissue from each individual by extraction with 80% methanol (MeOH) according to the protocol by Lee et al. (1987). First, HP was homogenized using an Ultra Turraxhomogenizer for one minute. 4 mL of 80% MeOH was added to 1g of homogenate, which was thoroughly mixed (Vortex minishaker) for two minutes. The mixture was centrifuged for 10 min (X g and the MeOH phase was collected and filtered through 0.22 [micro]m spin-X filters (Costar, Coming, NY, USA) before toxin analysis. Extraction of toxins from the algal samples was done by adding 10 mL of 80% MeOH to the filter papers and then ulstrasonicating the sample for 30 s. The extract volume was reduced by evaporation under [N.sub.2(g]) and then it was redissolved in 500 [micro]L 100% MeOH and filtered through a spin-X filter prior to analysis.

LC-MS/MS Analysis

The analysis of OA, DTX-1, DTX-2, PTX2, PTX2 SA, PTX12, and "PTX12 SA" were done using HPLC-MS/MS. Analytes were separated on a 3.0 x 100 mm Waters Symmetry C18 analytical column (3.5-[micro]m particles) using a gradient from 35% to 98% acetonitrile (MeCN) over 10 min, followed by an isocratic period of 6.5 min at 98% MeCN and an equilibration period of 3.5 min at 35% MeCN. The flow rate was 0.3 mL [min.sup.-1] and 2 mM ammonium formate and 50 mM formic acid was added to the mobile phases. Mass spectrometry was conducted on a Micromass QuattroUltima triple quadrupole mass spectrometer (Waters Micromass, Manchester, UK). For determination of OA-group toxins, MRM-chromatograms were recorded in negative ionization mode for the transitions m/z 803.5 [right arrow] 255.5 (OA/DTX-2) and m/z 817.5 [right arrow] 255.5 (DTX-1), and calibration curves were constructed for all of the three parent compounds. For quantitative analysis of the parent compounds, 1 mL of the shellfish extract was mixed with 410 [micro]L of water. For quantitative estimates of esters of the same three toxins, a hydrolysis step was performed prior to HPLC-MS/MS analysis. Two hundred micro liters of IM NaOH dissolved in 80% MeOH was added to 1 mL shellfish extract and the mixture was left to react at 37[degrees]C for 45 min. To stop the reaction and adjust the pH, 210 [micro]L of 1M H[Cl.sub.(aq)] were added. For determination of PTXs, MRM-chromatograms were recorded in positive ionization mode for the transitions m/z 876.5 [right arrow] 823.5 (PTX2), m/z 894.5 [right arrow] 823.5 (PTX2 SA), m/z 874.5 [right arrow] 821.5, and m/z 892.5 [right arrow] 821.5 ('PTX12 SA'). Calibration curves were constructed for PTX2 and PTX2 SA. The amount PTX12 was calculated from the PTX2 calibration curve, "PTX12 SA" was calculated from the PTX2 SA calibration curve, assuming equal responses on the MS-detector was made.

Total toxin content (ng [HP.sup.-1]), calculated as [toxic compound]*weight of HP, was used to compare accumulation and detoxification of the various DST compounds between species. By using toxin content in HP as a variable instead of toxin concentration, the HP size differences between the species are taken into account, as well as temporal changes in HP weights, which can affect the toxin concentration even if the toxin content remains unchanged in the tissue. The sum of all toxic components within each group of toxins was also calculated to compare data with the regulatory limit within the EU (160 [micro]g [kg.sup.-1] shellfish meat)

Statistical Analysis, Calculation of Reduction Rates

Differences between mussels and oysters in content of each OA-group parent compound, their esters and the various PTX compounds were analyzed using a two-factor analysis of variance (ANOVA) with species and weeks as fixed factors. For significant results (P < 0.05), Student-Neuman-Keuls (SNK) a posteriori test for differences among means was used.

Rates of reduction for each toxin compound were calculated using log-transformed data for toxin content (ng [HP.sup.-1]) and fitting a linear regression line to the data. The slope of the regression line was then used to calculate the reduction rate (depuration half-lives, [t.sub.1/2]) using the formula: [t.sub.1/2] = (log 0.5)/k, where k is the slope coefficient.


Occurrence of Dinophysis and Toxin Levels in Plankton Samples

A bloom situation of Dinophysis spp. was observed during the field exposure period (September 3 to October 1) in Bjornsund (Table 1). D. acuta dominated the Dinophysis community during the whole period, where this species constituted 58% to 76% of the total abundance of Dinophysis spp.. The second most abundant species was D. dens (14% to 28%), which is regarded as a sexual stage during the life cycle of D. acuta and hence probably belong to the same species (Reguera et al. 1995). D. acuta and D. dens together made up 78% to 95 % of the Dinophysis bloom. Other potentially toxic Dinophysis species were D. acuminata (2% to 11%), D. norvegica (1% to 13%) and D. rotundata (1% to 2%). Max abundance of all species was detected September 10, when the concentration of D. acuta exceeded 2,000 cells L-1. Analysis of the toxin profiles in plankton samples showed that the Dinophysis community contained toxins of both the OA and PTX-group (Table 2). The following toxins were detected: OA, DTX1, DTX2, PTX2, PTX2 SA, PTX12-1, PTX12-2, and "PTX12 SA." The seco acids (SA) of PTX2 and PTX12 do not occur naturally in the algae but can be formed during sample preparation via enzymatic hydrolysis. Therefore, PTX2 SA and PTX12 SA were combined with the nonhydrolyzed forms to calculate total PTX content in the samples. Also, PTX12-1 and PTX12-2 were combined and summed together. Overall, the PTX-group toxins dominated over the OA-group in the algae, indicated by a OA/ PTX ratio >1 (0.37-0.54, Table 2). Within the OA group, DTX-1 was the most abundant toxin detected, except on the first sampling occasion, when OA was the major toxin found. For the PTX-group, PTX2 was more abundant compared with PTX 12. The highest toxicity in the water samples was recorded on September 10, which correlated to the highest Dinophysis spp. concentrations (Table 1).

Accumulation of OA-Group Toxins During Field Exposure

DTX-1, OA and DTX-2 together with their respective acylesters were detected in both mussels (Fig. 1a to c) and oysters (Fig. 2a to c) during the field stage of the experiment. In accordance with the findings in the algae, DTX-1 predominated in the shellfish, followed by OA and DTX-2.

In mussels, a gradual temporal increase in all OA-group toxins occurred during the field exposure, where the highest levels were recorded after 4 wk of exposure (week 5), when mean content [+ or -] SE for DTX1 were 8640 [+ or -] 1950 ng [HP.sup.-l] DTX1-esters: 1590 [+ or -] 650 ng [HP.sup.-1] OA: 3670 [+ or -] 860 ng [HP.sup.-1], OA-esters: 360 [+ or -] 85 ng [HP.sup.-1], DTX2:2970 [+ or -] 540 ng [HP.sup.-1], and DTX2-esters: 780 [+ or -] 290 ng [HP.sup.-1]. At all sampling occasions, the content of the free forms of the toxin dominated in mussels. The ester proportion ranged between 3% and 49% for OA, between 0% and 16% for DTX1, and 0% and 21% for DTX2. Esters of DTX1 and DTX2 were only detected in a few of the mussels whereas esters of OA were found in all mussel samples.



In oysters, the accumulation pattern of OA, DTX1, and DTX2 during the field exposure stage was different compared with mussels (Fig. 2a to c). All OA-group components increased after one week of exposure compared with the start value, but no further accumulation in time was detected except for DTX1, which increased further on week 5 compared with previous sampling dates. For DTX2, a trend towards a reduction in content after the first week in the field was even seen. The highest detected level of OA was 30 [+ or -] 6 ng [HP.sup.-1] on week 5 and 200 [+ or -] 90 ng [HP.sup.-1] of OA-esters on week 3. Max content of DXT1 and DTX1-esters was 60 [+ or -] 12 and 180 [+ or -] 30 ng [HP.sup.-1] respectively on week 5. For DTX2 compounds, the highest level of the free form was found on week 5 (13 [+ or -] 10 ng [HP.sup.-1]) whereas DTX2-esters peaked on week 2 (70 [+ or -] 40 ng [HP.sup.-1]). The acyl-esters dominated in oysters, ranging between 67% and 95% for esters of OA, the same for DTXI was 68% and 95% and for DTX2, the ester proportion was 47% to 99%. Esters of OA-group toxins were detected in all oyster samples during the field exposure.


The statistical analysis showed that the content of OA, DTXI, and DTX2 was significantly higher in mussels compared with oysters at all sampling occasions (P < 0.001), where levels were about 10 50 times greater in mussels depending on sampling date and toxin component. The sum of OA-group toxins was recalculated as [micro]g OA-equivalents [kg.sub.-1] shellfish meat to compare the accumulated levels to the regulation limit for harvest (160 [micro]g OA-equivalents [kg.sub.-1] shellfish meat) (Fig. 3a). Already after one week in the field, the concentration in mussels was above this limit. After 4 wk of exposure (week 5), the level was about 10 times greater than the established safety limit in mussels. In oysters, the level of OA-group toxins never reached this limit during the exposure (Fig. 3a).

Accumulation of PTX-Group Toxins During Field Exposure

PTX2, PTX2 SA, PTX12-1, PTX12-2, and PTXI2 SA accumulated in both mussels and oysters during the field exposure (Table 3). PTX12-1 and PTX12-2 content was summed and are shown as a common mean (PTX12) in Table 3. The major PTX component detected was PTX2 SA in both species, with levels generally about 10 times greater than the other forms of PTX. A significant increase in PTX2 SA compared with previous samplings occurred on October l after 4 wk of field exposure in both species (mussels: 6716 [+ or -] 1198 ng [HP.sup.-1], oysters: 5430 [+ or -] 420 ng [HP.sup.-1]). In oysters, this temporal increase was also seen for the other PTX components whereas in mussels, levels of PTX12 and PTX12 SA reached a maximum already after two weeks of exposure. A species-specific comparison showed that the content of PTX12 was significantly lower (P < 0.05) in oysters compared with mussels at all sampling dates during the field exposure. For the other PTX components, there was no significant difference between species except October 1, when levels of both PTX2 and PTX12 SA was significantly higher (P < 0.05) in oysters compared with mussels. To summarize, data suggested differential assimilation patterns of the various PTX-group toxins in these species.

When summed and recalculated as lug PTX kg-l shellfish meat, levels were above the safety limit in both species at all sampling dates during field exposure (Fig. 3b). There was a trend towards generally higher concentrations in mussels, however, the difference between species were not statistically significant (P > 0.05).

Rates of Reduction for OA-Group Toxins

After 4 wk of exposure to Dinophysis spp. in the field (week 5 in the illustrations), mussels and oysters were transferred to algal-free seawater for detoxification in the laboratory. Detoxification was conducted during 7 wk and the results are shown in the right hand part in Figure 1 a-c and 2 a-c (Note: data for week 6 are missing because of an unfortunate mix-up of samples during analysis). Using data from week 5 as start value for detoxification, the rate of reduction for each component was calculated and the resulting half-lives ([t.sub.1/2]) in mussels and oysters are shown in Table 4. Because esters of DTX1 and DTX2 were only detected on a few occasions in mussels during detoxification, it was not possible to achieve a regression line and hence calculate [t.sub.1/2] for these compounds. Regarding the free forms of OA-group toxins in oysters, these were close to the detection limit or zero in some cases, hence the detoxification slopes were approximated from less data points in oysters compared with the mussels. [T.sub.1/2] was lowest for OA in both species compared with DTX1 and DTX2. The reduction rate of free OA was in the same range in both mussels ([t.sub.1/2] = 17 days) and oysters ([t.sub.1/2]=15 days) whereas reduction of DTX1 seemed to be slightly faster in mussels ([t.sub.1/2] = 23 days) compared with oysters ([t.sub.1/2] = 31 days). For free DTX2, the pattern was opposite, with [t.sub.1/2] being slightly lower in oysters ([t.sub.1/2] = 28 days) compared with mussels ([t.sub.1/2] = 33 days). For esters of OA, reduction rate was slower compared with free OA in both species, but OA-esters were reduced with a faster rate in oysters ([t.sub.1/2] = 23 days) compared with mussels ([t.sub.1/2] = 35 days). In oysters, esters of both DTX1 and DTX2 had a lower reduction rate ([t.sub.1/2] = 42 and 32 days respectively) compared with the OA-esters.

Although the concentration of all OA-group toxins was significantly reduced in both mussels and oysters after 7 wk of detoxification (week 12), total OA-group toxin level in mussels was still above the regulation limit when the experiment finished (Fig. 3a). The calculated half-life for the sum of all OA-group components was 24 days for mussels and 27 days for oysters (Table 4).

Rates of Reduction for PTX

A rapid reduction of all PTX compounds was seen already after one week of detoxification (October 8) in both species, when 80% to 90% of the various forms of PTX were lost (Table 3). The calculated half-lives of PTX2 and PTX2 SA were highly similar for both species ([t.sub.1/2] = 7-8 days, Table 4). For PTX12 and PTX12 SA, the rate of reduction was slightly slower compared with the PTX2 forms ([t.sub.1/2] = 11-13 days) except for PTXI2 in oysters. ([t.sub.1/2] = 6 days). The total PTX level was reduced below the regulation limit after 1 wk of detoxification in both species (Fig. 3b).


The main objective of this study was to compare the accumulation and detoxification of toxins originating from Dinophysis in two commercially important bivalve species and to test the general prediction that oysters accumulate toxins to a lesser extent than mussels. This hypothesis was derived from the results of a previous field study when no OA, DTX1, or DTX2 were detected in oysters during a DSP event in mussels (Svensson et al. 2000). The results strongly supported the previous observations, because the content of OA, DTX1, and DTX2 was 10-50 times greater in mussels compared with oysters depending on the sampling date during the four weeks of field exposure. The regulation limit for OA-group toxins was never exceeded in oysters in this experiment when at the same time, mussel toxicity reached a level 10 times greater than this limit. From the industrial perspective, this is important information, considering that the results were obtained during a moderate to severe bloom situation in terms of duration and intensity. The concentration of D. acuta, which is the most toxic Dinophysis species in Swedish and Norwegian waters (Dahl & Johannessen 2001, Lindahl et al. 2007), varied between 500-2038 cells [L.sup.-1] during the whole experimental period, which is several times greater than the advisory limit of 200 cells [L.sup.-1] of D. acuta set by the food safety authorities in Sweden and Norway for an increased risk of accumulation of DST in mussels. Regarding the toxin profiles in the plankton samples, DTX1 was the major OA-group toxin detected, which reflected the dominance of D. acuta in the Dinophysis spp. community. This species has previously been confirmed to be the major producer of DTX1 in this fjord area (Lindahl et al. 2007), DTX1 was also the main OA-group parent compound detected in the shellfish during the experiment.

As far as we know, a comparison between the accumulation and detoxification of OA-group toxins has not been done previously in these species. Some field data on DSP toxins in Mytilus sp. and the oyster Crassostrea gigas suggests that C. gigas do not accumulate or accumulate at least 10 times less compared with the mussels (Poletti et al. 1998, Vale & Sampayo 2002a). Other data collected in the field have shown that Mytilus spp. generally contain the highest total levels of these toxins compared with various species of cockles, scallops, and clams (Vale & Sampayo 2002a, Vale & Sampayo 2002b, Vale 2004, Villar-Gonzalez et al. 2007, Suzuki & Mitsuya 2001). Together with the results shown in this paper, further support for Mytilus spp. being a top candidate among bivalve species to accumulate DSP toxins is given.

Our second prediction was that the ester forms of OA-group toxins would dominate in the oysters if detected. This prediction was also supported by the results in this study, because OA-group toxins were mainly in the form of esters in oysters, ranging from 67% to 99% of the total toxin content. In mussels, esters of OA were detected in all samples but never exceeded 49% of total toxin content. Esters of DTX1 and DTX2 acylesters were only detected occasionally in mussels and the proportion was lower compared with OA-esters, ranging from 0% to 16% for DTX1 and 0% to 21% for DTX2. The ester proportion in Mytilus spp. has previously been shown to be consistently lower compared with other bivalve species such as various clams (Spisula spp., Ensis spp, Ruditapes decussate, Solen marginatus), peppery furrow shell (Scobicularia phma), carpet shell (Venerupis vellastra), cockles (Cerastoderma edule), scallops (Patinopecten yessoensis), and oysters (Crassostrea japonica) (Jorgensen et al. 2005, Suzuki & Mitsuya 2001, Vale & Sampayo 2002a, Vale & Sampayo 2002b, Vale 2004, Vale 2006, Villar-Gonzalez et al. 2007). One exception is M. chilensis, where the esters of DTX1 have been found to constitute 98% to 100% of total toxin burden (Garcia et al. 2004, Garcia et al. 2005, Garcia et al. 2006). Caution regarding conclusions about the absolute amounts of acyl-esters should be used, because recent evidence suggests that tissue extraction in 80% MetOH, initially recommended by Lee et al. (1987) for extraction of OA and which was used in this study, is a poor extraction solvent for the more lipid-soluble esters (Jorgensen et al. 2005, McNabb et al. 2005). Instead, repeated extractions using 90% to 100% MeOH have been found to be the most efficient solvent for DSP esters in shellfish tissue. Compared with using 80% MeOH, which recovers between 64% and 80% of the esters, the use of 90% MeOH recovers 93% to 98% (Jorgensen et al. 2005). Hence, the ester content and proportion is likely to be underestimated in oysters and mussels in this study, which needs to be taken into consideration when the results are interpreted. As discussed in Jorgensen et al. (2005), many previous studies which have used 80% MetOH for sample preparation must be expected to have reported too low DSP ester contents and ester percentage. Nevertheless, underestimation of the ester content is not likely to have affected the relative difference in toxin content between oysters and mussels in this study, because all steps in the sample preparation were equally performed. The exception could be if mussels and oysters display large differences in fatty acid profiles (i.e., chain lengths and degree of saturation) of the esters, because esters with saturated and/or long-chained fatty acid moieties will have lower solubility in 80% MeOH compared with esters of shorter and/or unsaturated fatty acids. However, direct analysis of the fatty acid profiles of the OA-group toxins showed a high similarity in the distribution of esters in oysters and mussels during this experiment (Torgersen et al. 2008). The profiles were in accordance with the free fatty acid profiles in oysters and mussels, dominated by palmitic acid (16:0). This result suggests that the conclusions about strong species-specific differences in accumulation of OA-group toxins are valid. When levels and dynamics of the PTX-group toxins were compared between oysters and mussels, differences were less apparent. For the flat oyster, this is the first time uptake of PTX is documented, whereas in Mytilus spp., accumulation of various PTX compounds has been demonstrated on several occasions. Both mussels and oysters rapidly accumulated levels of PTX compounds above the regulation limit already after one week of exposure. As for the OA-group toxins, oysters generally contained lower total amounts of PTX compared with mussels, but the difference was nonsignificant and after four weeks of exposure, PTX levels were in the same range in both species. Some differential assimilation patterns of the various PTX-group toxins were seen but both species contained PTX2 SA as the major PTX compound, being about 10 times greater than the other forms of PTX. This pattern is in line with results generally seen in most shellfish species, where PTX2 is rapidly hydrolyzed to PTX2 SA in the bivalve tissue (Daiguji et al. 1998, Suzuki et al. 2001a, Suzuki et al. 2001b, MacKenzie et al. 2002, Miles et al. 2004, Vale 2004, Blanco et al. 2007, Villar-Gonzalez et al. 2007).

PTX2 SA was previously believed to be the major metabolic product of PTX2 in shellfish. Because the PTX compounds are destroyed during alkaline hydrolysis, which is the common method to detect esters of OA-group toxins, esters of PTX were probably not seen before in shellfish samples. Wilkins et al. (2006) identified a series of fatty acid monoester derivatives of PTX2 SA when screening for PTX derivatives in unhydrolyzed extracts from blue mussels. The amount ofPTX2 SA esters were more than 20-fold higher than the amount of unesterified PTX2 SA, similar to what previously been seen for the ratio of esterified to unesterified OA-group toxins in scallops, cockles, clams, and now also oysters O. edulis. Analysis of PTX2 SA ester profiles in mussels and oysters from this experiment showed a more diverse range of fatty acid side chains of PTX2 SA compared with the OA-group toxins and also differences between mussels and oysters in major fatty acids of PTX2 SA (Torgersen et al. 2008). PTX2 SA was esterified to a larger degree in mussels (81%) compared with oysters (64%), which was opposite to the esterification of OA-group toxins. When all PTX2 SA esters were summed, mussels contained about 5-fold higher levels of PTX2 SA esters (28 600 ng [HP.sup.-1], Torgersen et al. 2008) compared with unesterified PTX2 SA (6,716 ng [HP.sup.-1], Table 3), whereas esters of PTX2 SA in oysters were only about 2-fold higher (10 500 ng [HP.sup.-1], Torgersen et al. 2008) compared with PTX2 SA (5,430 ng [HP.sup.-1], Table 3). The relative difference between mussels and oysters in accumulation of PTX consequently becomes larger if PTX2 SA esters are included in the PTX budget. The findings by Wilkins et al. (2006) and Torgersen et al. (2008), that PTX2 SA esters are highly abundant in blue mussels and flat oysters, imply that this is likely to be the case for other shellfish species too, which needs to be considered in the interpretation of previous research and when planning future studies concerning PTX.

When mussels and oysters were transferred to clean water for detoxification, all toxins were greatly reduced after 7 wk of detoxification in both species. [T.sup.1/2] was calculated for each toxin separately and for the sum of all toxins to evaluate whether faster reduction rates could explain the lower toxicity in oysters. For the OA-group toxins, there was, however, no consistent species-specific difference in reduction half-lives that would suggest that oysters eliminate the OA-group compounds faster compared with mussels. The only exception would be esters of OA, where the reduction rate was 23 days in oysters compared with 35 days in mussels. Elimination of DTX1 and DTX2 esters could not be computed in mussels because these were only detected occasionally by the indirect hydrolysis method. However, data from Torgersen et al. (2008), who used direct detection of esters in the shellfish extracts, showed that [t.sub.1/2] for esters of DTX1 and DTX2, respectively, were very similar in mussels and oysters. Torgersen et al. (2008) also showed that the elimination of DTX1 esters was significantly slower in both species compared with esters of OA and DTX2. Compared with the esterified forms, our data indicated that the free forms of OA, DTX1, and DTX2 was reduced at a faster rate than their respective esters in both species, a pattern which was particularly evident for OA in mussels ([t.sub.1/2] 17 days for OA, [t.sub.1/2] = 35 days for OA-esters). This is in contrast to results found by Vale (2004), where esters of OA and DTX2 were eliminated faster compared with their free forms in mussels (M. galloprovincialis) and cockles (C. edule). Other studies have indicated the opposite, for example, slower reduction of the esters of DST in M. galloprovincialis (Fernandez et al. 1998, Morono et al. 2003). The calculated half-life of all OA-compounds summed together was 24 days in mussels and 27 days in oysters. The PTX group toxins were reduced at a considerably faster, but equal, rate in both species compared with the OA-group toxins. [T.sub.1/2] was 8 days in mussels and 9 days in oysters for the sum of all PTX. Only PTX12, found in low concentrations in oysters, appeared to be eliminated at a faster rate in oysters ([t.sub.1/2] = 6 days) compared with mussels ([t.sub.1/2] = 13 days). Vale (2004) also confirmed rapid reduction of PTX2 and PTX2 SA in M. galloprovincialis during depuration. Slower reduction rate of PTX2 SA ([t.sub.1/2] = 26 days) was reported in Greenshell mussels (Perna canaliculus) from New Zealand (MacKenzie et al. 2002). To summarize, our results did not provide convincing evidence for faster elimination rates of OA and PTX toxin groups in oysters, which would explain the large differences in retained levels of toxins between mussels and oysters. Instead, differential rates of gut assimilation and/or biotransformation of the OA and PTX-group toxins may explain some of the observed differences in retention and toxin profiles between the bivalves. To quantitatively estimate the assimilation and biotransformation rates of various toxins, controlled in vivo or in vitro experiments using purified toxins need be performed. Also, models related to differences in feeding rates, particle selection and behavioural response to toxic algae can be proposed to explain the lower accumulation of toxins in oysters. Different bivalve species respond differently to toxigenic algae, which can be seen as reduced clearance, feeding inhibition and/or particle selection, and rejection. There are several examples of a negative feeding response to various toxin-producing algae in oyster species (Cassis & Taylor 2004, Gainey & Shumway 1988, Shumway et al. 2006) but also in other bivalves such as hard clams (Mercenaria mercenaria) and M. edulis (Bricelj et al. 2004, Shumway et al. 2006) and soft clams (Mya arenaria) (Bricelj et al. 2005). These preingestive theories need to be tested specifically for Dinophysis spp. in comparative experiments including both bivalve species.


This study has provided strong support for interspecies differences in accumulation of toxins belonging to the DSP complex between two commercially important bivalve species. From the industrial perspective, the fact that the toxicity never exceeded the regulatory limit is important information in particular because the oysters were exposed to a dense and extended bloom of Dinophysis spp. Communication with the Danish, Norwegian, and Irish food authorities confirm that toxins of the OA-group are seldom detected in O. edulis. Flat oysters should thus be regarded as a low-risk species for OA-group toxin contamination, which should be taken into consideration by the regulatory authorities in charge of sampling frequencies and monitoring programs of shellfish toxins. Concerning the PTX compounds, the oral toxicity and hence the risk to human health of particularly the PTX2 SA, which dominated in both species, is debated. PTX2 is highly toxic when injected intraperitoneally (i.p.) into mice, whereas PTX2 SA does not cause any i.p. effects (Miles et al. 2004). When administered by the oral route, neither compounds cause diarrhea or other effects in mice at doses up to 5,000 [micro]g [kg.sub.-1] (Miles et al. 2004). Because oysters and mussels almost exclusively contain PTX SA and esters thereof, health problems caused by PTX are not expected in commercially exploited bivalves in Scandinavia.


This work has been supported by grants to S. Lindegarth from the Swedish Fisheries Board (EU EFF fundings), Stiftelsen Magnus Bergwall and Makarna Wahlstroms minnesfond. Also, the Research Council of Norway (grant no. 164851S40) is gratefully acknowledged for financial support to T. Torgersen.


Blanco, J., G. Alvarez & E. Uribe. 2007. Identification of pectenotoxins in plankton, filter-feeders, and isolated cells of a Dinophysis acuminata with an atypical toxin profile, from Chile. Toxicon 49:710-716.

Bricelj, M. V., S. P. MacQuarrie & R. Smolowitz. 2004. Concentration-dependent effects of toxic and non-toxic isolates of the brown tide alga Auerococcus anophagefferens on growth of juvenile bivalves. Mar. Ecol. Prog. Ser. 282:101-114.

Bricelj, M. V., L. Connel, K. Konoki, S. P. MacQuarrie, T. Scheuer, W. A. Catterall & V. L. Trainer. 2005. Sodium channel mutation leading to saxitoxin resistance in clams increase risk of PSP. Nature 434:763-767.

Cassis, D. & F. J. R. Taylor. 2004. Rapid responses of juvenile oysters exposed to potentially harmful phytoplankton. 5th Int. Conf. on Molluscan Shellfish Safety, Galway, Ireland. pp. 170-174.

Dahl, E. & T. Johannessen. 2001. Relationship between occurrence of Dinophysis species (Dinophyceae) and shellfish toxicity. Phycologia 40:223-227.

Daiguji, M., M. Satake, K. J. James, A. Bishop, A. L. MacKenzie, H. Naoki & T. Yasumoto. 1998. Structures of new pectenotoxin analogs, pectenotoxin-2 seco acid and 7-epi-pectenotoxin-2 seco acid, isolated from a dinoflagellate and Greenshell mussels. Chem. Lett. 27:653-654.

Daranas, A. H., M. Norte & J. J. Fernandez. 2001. Toxic marine microalgae. Toxicon 39:1101-1132.

Fernandez, M. L., A. Miguez, A. Morono, E. Cacho, A. Martinez & J. Blanco. (1998) Detoxification of low polarity toxins (DTX-3) from mussels Mytilus galloprovincialis in Spain. In: B. Reguera, J. Blanco, M. L. Fernandez & T. Wyatt, editors. Harmful algae. Xunta de Galicia and Intergovernmental Oceanographic Commission of UNESCO. pp. 449-452.

Garcia, C., V. Gonzalez, C. Cornejo, H. Palma-Fleming & N. Lagos. 2004. First evidence of Dinophysistoxin-1 ester and carcinogenic polycyclic aromatic hydrocarbons in smoked bivalves collected in the Patagonia fjords. Toxicon 43:121-131.

Garcia, C., D. Truan, M. Lagos, J. P. Santelices, J. C. Diaz & N. Lagos. 2005. Metabolic transformation of dinophysistoxin-3 into dinophysistoxin-1 causes human intoxication by consumption of O-acylderivatives dinophysistoxins contaminated shellfish. J. Toxicol. Sci. 30:287-296.

Garcia, C., D. Truan, M. Lagos, J. P. Santelices, J. C. Diaz & N. Lagos. 2006. High amounts of dinophysistoxin-3 in Mytilus chilensis collected in Seno de Reloncavi, Chile, during massive human intoxication associated with outbreak of Vibrio parahaemolyticus. J. Toxicol. Sci. 31:305-314.

Gainey, L. F. & S. E. Shumway. 1988. A compendium of the responses of bivalve molluscs to toxic dinoflagellates. J. Shellfish Res. 7:623-628.

Jorgensen, K., S. Scanlon & L. B. Jensen. 2005. Diarrhetic shellfish poisoning toxin esters in Danish blue mussels and surf clams. Food Addit. Contain. 22:743-751.

Karlson, B., A.-S. Rehnstam-Holm & L.-O. Loo. 2007. Temporal and spatial distribution of diarrhetic shellfish toxins in blue mussels, Mytilus edulis (L.), at the Swedish west coast, NE Atlantic, years 1988 to 2005. Swedish Meteorological and Hydrological Institute, Reports Oceanography No. 35. 40 pp.

Krogh, P., L. Edler, E. Graneli & U. Nyman. 1985. Outbreak of Diarrhetic Shellfish Poisoning on the west coast of Sweden. In: D. A. Anderson, A. W. White & D. G. Baden, editors. Toxic dinoflagellates. Elsevier Science Publishing, Amsterdam. pp. 501-503.

Lee, J. S., T. Yanagi, R. Kenma & T. Yasumoto. 1987. Fluorometric determination of diarrhetic shellfish toxins by high pressure liquid chromatography. Agric. Biol. Chem. 51:877-881.

Lindahl, O., B. Lundve & M. Johansen. 2007. Toxicity of Dinophysis spp. in relation to population densitiy and environmental conditions on the Swedish west coast. Harmful Algae 6:218-231.

MacKenzie, L., P. Holland, P. McNabb, V. Beuzenberg, A. Selwood & T. Suzuki. 2002. Complex toxin profiles in phytoplankton and greenshell mussels (Perna eanaliculus), revealed by LC-MS/MS analysis. Toxicon 40:1321-1330.

McNabb, P., A. I. Selwood, P. Holland, J. Aasen, T. Aune, G. Eaglesham, P. Hess, M. Igarishi, M. Quilliam, D. Slattery, J. Van de Riet, H. Van Egmond, H. Van den Top & T. Yasumoto. 2005. Multiresidue method for determination of algal toxins in shellfish: single-laboratory validation and interlaboratory study. J. AOAC Int. 88:761-772.

Miles, C. O., A. L. Wilkins, R. Munday, M. H. Dines, A. D. Hawkes, L. R. Briggs, M. Sandvik, D. J. Jensen, J. M. Cooney, P. T. Holland, M. A. Quilliam, A. L. MacKenzie, V. Beuzenberg & N. R. Towers. 2004. Isolation of pectenotoxin-2 from Dinophysis acuta and its conversion to pectenotoxin-2 seco acid, and preliminary assessment of their acute toxicities. Toxicon 43:1-9.

Morono, A., F. F. Arevalo, M. L. Fernandez, J. Maneiro, Y. Pazos, C. Salgado & J. Blanco. 2003. Accumulation and transformation of DSP toxins in mussels Mytilus galloprovincialis during a toxic episode caused by Dinophysis acuminata. Aquat. Toxicol. 62:269-280.

Poletti, R., K. Cettul, F. Bovo, A. Milandri, M. Pompei & R. Frate. 1998. Distribution of toxic dinoflagellates and their impact on shellfish along the northwest Adriatic coast. In: B. Reguera, J. Blanco, M. L. Fernandez & T. Wyatt, editors. Harmful algae. Xunta de Galicia and Intergovernmental Oceanographic Commission of UNESCO. pp. 88-90.

Reguera, B., I. Bravo & S. Fraga. 1995. Autoecology and some life history stages of Dinophysis acuta Ehrenberg. J. Plankton Res. 17:99-1015.

Shumway, S. E., J. M. Burkholder & J. Springer. 2006. Effects of the estuarine dinoflagellate Pfisteria shumwayae (Dinophyceae) on survival and grazing activity of several shellfish species. Harmful Algae 5:442-458.

Suzuki, T. & T. Mitsuya. 2001. Comparison of dinophysistoxin-1 and esterified dinophysistoxin-1 (dinophysistoxin-3) content in the scallop Pactinopecten yessoensis and the mussel Mytilus galloprovincialis. Toxicon 39:905-908.

Suzuki, T., L. MacKenzie, D. Stirling & J. Adamson. 2001a. Pectenotoxin-2 seco acid: a toxin converted from pectenotoxin-2 by the New Zealand Greenshell mussel, Perna canaliculus. Toxicon 39:507-514.

Suzuki, T., L. MacKenzie, D. Stirling & J. Adamson. 2001b. Conversion of pectenotoxin-2 to pectenotoxin-2 seco acid in the New Zealand scallop, Pecten novaezelandiae. Fish. Sci. 67:506-510.

Svensson, S., C. Andre, A.-S. Rehnstam-Holm & J. Hansson. 2000. A case of consistent spatial differences in content of Diarrhetic Shellfish Toxins (DST) among three bivalve species: Mytilus edulis, Ostrea edulis and Cerastoderma edule. J. Shellfish Res. 19:1017-1020.

Torgersen, T., M. Sandvik, B. Lundve & S. Lindegarth. 2008. Profiles and levels of okadaic acid group toxins and pectenotoxins during toxin depuration. Part II: blue mussels (Mytilus edulis) and flat oysters (Ostrea edulis). Toxieon 52:418-427.

Vale, P., Sampayo & M. A. M. 2002a. Esterification of DSP toxins by Portuguese bivalves from the Northwest coast determined by LC-MS-a widespread phenomenon. Toxicon 40:33-42.

Vale, P., Sampayo, M. A. M. 2002b. First confirmation of human diarrhoeic poisonings by okadaic acid esters after ingestion of razor clams (Solen marginatus) and green crabs (Carcinus maenas) in Aveiro lagoon, Portugal and detection of okadaic acid esters in phytoplankton. Toxicon 40:989-996.

Vale, P. 2004. Differential dynamics of dinophysistoxins and pectenotoxins between blue mussel and common cockle: a phenomenon originating from the complex toxin profile of Dinophysis acuta. Toxicon 44:123-134.

Vale, P. 2006. Detailed profiles of 7-O-acyl esters in plankton and shellfish from the Portuguese coast. J. Chromatogr. A 1128: 181-188.

Van Dolah, F. M. 2000. Marine algal toxins: origins, health effects, and their increased occurrence. Environ. Health Perspect. 108:133-141.

Villar-Gonzalez, A., M. L. Rodriguez-Velasco, B. Ben-Gigirey & L. M. Botana. 2007. Lipophilic toxin profile in Galicia (Spain): 2005 toxic episode. Toxicon 49:1129-1134.

Wilkins, A. L., N. Rehmann, T. Torgersen, T. Rundberget, M. Keogh, D. Petersen, P. Hess, F. Rise & C. O. Miles. 2006. Identification of fatty acid esters of pectenotoxin-2 seco acid in blue mussels (Mytilus edulis) from Ireland. J. Agric. Food Chem. 54:567-5678.

Yasumoto, T., Y. Oshima & M. Yamaguchi. 1978. Occurence of a new type of shellfish poisoning in the Tohoku district. Bull. Jpn. Soc. Sci. Fish. 44:1249-1255.

Yasumoto, T., M. Murata, J. S. Lee & K. Torigoe. 1989. Polyether toxins produced by dinoflagellates. In: S. Natori, K. Hashimoto & Y. Ueno, editors. Mycotoxins and phycotoxins'88. Amsterdam: Elsevier. pp. 375-382.


(1) Dept. of Marine Ecology-Tjarno, Goteborg University, S-45296 Stromstad, Sweden; (2) National Veterinary Institute, Dept. of Feed and Food Safety, P.O. Box 750 Sentrum, 0106 Oslo, Norway; (3) Dept. of Marine Ecology-Kristineberg, Goteborg University, Kristineberg 566, 450 34 Fiskebackskil, Sweden

* Corresponding author. E-mail:
Abundance (cells L-1) of Dinophysis sp. in Bjornsund during the
experimental period. The relative abundance (%) of each species
to the total Dinophysis community is shown within brackets.

           Week       D.          D.          D.
Date       no.      acuta      acuminata   norvegica

3rd Sep     1      960 (64)     115 (8)    203 (13)
10th Sep    2     2,038 (68)    142 (5)     257 (9)
17th Sep    3      737 (69)      34 (3)       7 (1)
24th Sep    4      514 (58)     95 (11)       7 (1)
1st Oct     5      500 (76)      14 (2)       7 (1)

               D.          D.
Date       rotundata     dens

3rd Sep      14 (1)    216 (14)
10th Sep     27 (1)    520 (17)
17th Sep     14 (1)    277 (26)
24th Sep     20 (2)    243 (28)
1st Oct      14 (2)    122 (19)

Concentrations of OA-groups toxins and PTX in plankton samples,
expressed as ng 100 [L.sup.-1] seawater, in Bjornsund during field
exposure (Sep 3 to Oct 1). Also, the calculated sum of OA-group toxins
and PTX and the ratio between OA and PTX is shown. Note that PTX-12 is
the sum of PTX12-1 and PTX12-2. Seco acids (SA) of both PTX-2 and
PTX-12 were also detected in minor concentrations in the samples,
these are included in the PTX-2 and PTX-12 data.

           Week                          Sum
Date       no.    OA    DTX-1   DTX-2    OA     PTX-2

3rd Sep     1     197    165       69     430     644
10th Sep    2     458    870      193   1,522   2,362
17th Sep    3     268    654      107   1,029   1,406
24th Sep    4      96    543       46     684   1,476
1st Oct     5      59    565       17     641   1,076

                    Sum     OA/
Date       PTX-12   PTX     PTX

3rd Sep     418     1,062   0.40
10th Sep    759     3,120   0.49
17th Sep    544     1,950   0.53
24th Sep    387     1,863   0.37
1st Oct     119     1,196   0.54

Content of PTX2, PTX2 SA, PTX12 (sum of PTX12a and PTX12b), and PTX12
SA, expressed as ng [HP.sup.-1], in mussels and oysters during field
exposure (Sep 3 to Oct 1) and laboratory detoxification (Start Oct 1
to Nov 18). Data are mean values [+ or -] SE, n = 5.

Date      Week no.   Species         PTX2               PTX2 SA

3rd Sep      1       Mussel           0               5 [+ or -] 1
                     Oyster           0              99 [+ or -] 35
10th Sep     2       Mussel    215 [+ or -] 58    2,685 [+ or -] 911
                     Oyster    139 [+ or -] 50    1,539 [+ or -] 534
17th Sep     3       Mussel    183 [+ or -] 58    2,467 [+ or -] 1226
                     Oyster    101 [+ or -] 41    1,227 [+ or -] 462
24th Sep     4       Mussel    237 [+ or -] 62    2,497 [+ or -] 844
                     Oyster    141 [+ or -] 53    1,459 [+ or -] 488
1st Oct      5       Mussel    234 [+ or -] 42    6,716 [+ or -] 1198
                     Oyster    556 [+ or -] 111   5,430 [+ or -] 420
8th Oct      6       Mussel     29 [+ or -] 6     1,042 [+ or -] 293
                     Oyster      7 [+ or -] 1       201 [+ or -] 38
15th Oct     7       Mussel     10 [+ or -] 2       164 [+ or -] 74
                     Oyster      2 [+ or -] 1        31 [+ or -] 13
22nd Oct     8       Mussel      9 [+ or -] 3       171 [+ or -] 47
                     Oyster      5 [+ or -] 2        65 [+ or -] 16
29th Oct     9       Mussel      5 [+ or -] 1       123 [+ or -] 29
                     Oyster      1 [+ or -] 0       28 [+ or -] 13
5th Nov      10      Mussel      4 [+ or -] 2       84 [- or -] 19
                     Oyster      1 [+ or -] 0       35 [+ or -] 13
12th Nov     11      Mussel      2 [+ or -] 0       43 [+ or -] 14
                     Oyster      1 [+ or -] 1       28 [+ or -] 9
18th Nov     12      Mussel      1 [+ or -] 0       32 [+ or -] 7

Date              PTX12            PTX12 SA

3rd Sep             0                 0
                    0                 9
10th Sep     306 [+ or -] 69   161 [+ or -] 50
              36 [+ or -] 15   269 [+ or -] 96
17th Sep     377 [+ or -] 87   347 [+ or -] 117
              78 [+ or -] 34   384 [+ or -] 146
24th Sep     305 [+ or -] 68   251 [+ or -] 63
              36 [+ or -] 15   200 [+ or -] 55
1st Oct      143 [+ or -] 26   187 [+ or -] 19
              96 [+ or -] 8    592 [+ or -] 68
8th Oct       21 [+ or -] 6     33 [+ or -] 8
               1 [+ or -] 1     60 [+ or -] 17
15th Oct      17 [+ or -] 3     17 [+ or -] 5
               1 [+ or -] 0     10 [+ or -] 4
22nd Oct      22 [+ or -] 6     18 [+ or -] 4
                    0           40 [+ or -] 13
29th Oct      12 [+ or -] 1     16 [+ or -] 2
                    0           16 [+ or -] 9
5th Nov       13 [+ or -] 3     17 [+ or -] 4
                    0           16 [+ or -] 7
12th Nov       6 [+ or -] 1      7 [+ or -] 2
                    0           10 [+ or -] 4
18th Nov       5 [+ or -] 1      6 [+ or -] 1
                    0           11 [+ or -] 2

Table 4.
Calculated half-lives (TI/2) of the detected toxin compounds
during the laboratory detoxification stage of the experiment
(Oct 1 to Nov 18). n.a. = not applicable (too few data points).

Toxin          Species   [T.sub.1/2] (days)

OA             Mussel             17
               Oyster             15
OA esters      Mussel             35
               Oyster             23
DTX-1          Mussel             23
               Oyster             31
DTX-1 esters   Mussel           n.a.
               Oyster             42
DTX-2          Mussel             33
               Oyster             28
DTX-2 esters   Mussel            n.a
               Oyster             32
PTX2           Mussel              8
               Oyster              8
PTX2 SA        Mussel              7
               Oyster              8
PTX 12         Mussel             13
               Oyster              6
PTX12 SA       Mussel             12
               Oyster             11
sum of DST     Mussel             24
               Oyster             27
sum of PTX     Mussel              8
               Oyster              9
COPYRIGHT 2009 National Shellfisheries Association, Inc.
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2009 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Author:Lindegarth, Susanne; Torgersen, Trine; Lundve, Bengt; Sandvik, Morten
Publication:Journal of Shellfish Research
Article Type:Report
Geographic Code:1USA
Date:Apr 1, 2009
Previous Article:Energy storage and reproduction in mussels, Mytilus galloprovincialis: the influence of diet quality.
Next Article:Non-destructive method to study the internal anatomy of the Chilean scallop Argopecten purpuratus.

Related Articles
Antimicrobial activity of copper and zinc accumulated in eastern oyster amebocytes.
Effects of polynuclear aromatic hydrocarbons on hemocyte characteristics of the Pacific oyster, Crassostrea gigas.
Morphological, structural and functional characteristics of the hemocytes of the oyster, Crassostrea ariakensis.
Diseases of pearl oysters and other molluscs: a Western Australian perspective.
The susceptibility of young prespawning oysters, Ostrea edulis, to Bonamia ostreae.
Genetic diversity of the European oyster (Ostrea edulis L.) in Nova Scotia: comparison with other parts of Canada, Maine and Europe and implications...
Spat collection of a non-native bivalve species (European oyster, Ostrea edulis) off the Eastern Canadian Coast.
Susceptibility of Crassostrea ariakensis (Fujita 1913) to Bonamia and Perkinsus spp. infections: potential for disease transmission between oyster...
Larval quality of a nonnative bivalve species (European oyster, Ostrea edulis) off the east Canadian coast.
Nitric oxide and superoxide anion generation in hemocytes of Crassostrea ariakensis stimulated with rickettsia-like organisms and zymosan.

Terms of use | Privacy policy | Copyright © 2021 Farlex, Inc. | Feedback | For webmasters