Effects of Heating on Proportions of Azaspiracids 1–10 in Mussels (Mytilus edulis) and Identification of Carboxylated Precursors for Azaspiracids 5, 10, 13, and 15
Abstract:Access and use of this website and the material on it are subject to the Terms and Conditions set forth at Effects of heating on proportions of azaspiracids 1−10 in mussels (Mytilus edulis) and identification of carboxylated precursors for azaspiracid-5, -10, -13 and -15 Kilcoyne, Jane; McCarron, Pearse; Hess, Philipp; Miles, Christopher O.http://nparc.cisti-icist.nrc-cnrc.gc.ca/npsi/jsp/nparc_cp.jsp?lang=fr L'accès à ce site Web et l'utilisation de son contenu sont assujettis aux conditions présentées dans le… Show more
“…Mussels collected from Bruckless, Ireland, in 2005 have previously been used for the isolation of AZA analogues, and for the production of reference materials ( Hess et al, 2007 ; Kilcoyne et al, 2015a , 2015b ; McCarron et al, 2015 ). Given that AZAs are sensitive to the base hydrolysis conditions typically used for indirect analysis of fatty acid esters ( Alfonso et al, 2008 ), the Bruckless mussels were evaluated for intact fatty acid esters.…”
Section: Resultsmentioning
confidence: 99%
“…Two of the regulated analogues, AZA1 and -2, are produced by A. spinosum , while many others are shellfish metabolites and products from oxidation, hydroxylation, decarboxylation and dehydration ( Fig. 1 ) ( Hess et al, 2014 ; Kilcoyne et al, 2015a , 2018 ; McCarron et al, 2009 ; Tillmann et al, 2009 ). Fig.…”
Azaspiracids (AZAs) are lipophilic polyether toxins produced by
Azadinium
and
Amphidoma
species of marine microalgae. The main dinoflagellate precursors AZA1 and AZA2 are metabolized by shellfish to produce an array of AZA analogues. Many marine toxins undergo fatty acid esterification in shellfish, therefore mussel tissues contaminated with AZAs were screened for intact fatty acid esters of AZAs using liquid chromatography-high resolution mass spectrometry. Acyl esters were primarily observed for AZAs containing hydroxy groups at C-3 with 3-
O
-palmitoylAZA4 identified as the most abundant acyl ester, while other fatty acid esters including 18:1, 16:1, 17:0, 20:2 and 18:0 acyl esters were detected. The structures of these acyl derivatives were determined through LC-MS/MS experiments, and supported by periodate cleavage reactions and semi-synthesis of palmitate esters of the AZAs. Esters of the hydroxy groups at C-20 or C-21 were not observed in mussel tissue. The relative proportion of the most abundant AZA ester was less than 3% of the sum of the major free AZA analogues. These findings reveal an additional metabolic pathway for AZAs in shellfish.
“…Mussels collected from Bruckless, Ireland, in 2005 have previously been used for the isolation of AZA analogues, and for the production of reference materials ( Hess et al, 2007 ; Kilcoyne et al, 2015a , 2015b ; McCarron et al, 2015 ). Given that AZAs are sensitive to the base hydrolysis conditions typically used for indirect analysis of fatty acid esters ( Alfonso et al, 2008 ), the Bruckless mussels were evaluated for intact fatty acid esters.…”
Section: Resultsmentioning
confidence: 99%
“…Two of the regulated analogues, AZA1 and -2, are produced by A. spinosum , while many others are shellfish metabolites and products from oxidation, hydroxylation, decarboxylation and dehydration ( Fig. 1 ) ( Hess et al, 2014 ; Kilcoyne et al, 2015a , 2018 ; McCarron et al, 2009 ; Tillmann et al, 2009 ). Fig.…”
Azaspiracids (AZAs) are lipophilic polyether toxins produced by
Azadinium
and
Amphidoma
species of marine microalgae. The main dinoflagellate precursors AZA1 and AZA2 are metabolized by shellfish to produce an array of AZA analogues. Many marine toxins undergo fatty acid esterification in shellfish, therefore mussel tissues contaminated with AZAs were screened for intact fatty acid esters of AZAs using liquid chromatography-high resolution mass spectrometry. Acyl esters were primarily observed for AZAs containing hydroxy groups at C-3 with 3-
O
-palmitoylAZA4 identified as the most abundant acyl ester, while other fatty acid esters including 18:1, 16:1, 17:0, 20:2 and 18:0 acyl esters were detected. The structures of these acyl derivatives were determined through LC-MS/MS experiments, and supported by periodate cleavage reactions and semi-synthesis of palmitate esters of the AZAs. Esters of the hydroxy groups at C-20 or C-21 were not observed in mussel tissue. The relative proportion of the most abundant AZA ester was less than 3% of the sum of the major free AZA analogues. These findings reveal an additional metabolic pathway for AZAs in shellfish.
“…The pellets were further extracted using an Ultra Turrax for 1 min with an additional 9 mL of MeOH, centrifuged at 3950 g (5 min), and the supernatants decanted into the same 25-mL volumetric flasks, which were brought to volume with MeOH. A portion (10 mL) of each extract was transferred into sealed centrifuge tubes and placed for 10 min in a water bath heated to 90 °C to decarboxylate carboxylated AZAs [31,32]. The raw and heat-treated samples were then passed through Whatman 0.2-µm cellulose acetate filters into HPLC vials for analysis.…”
Section: Raw and Heat-treated Mussel Tissuesmentioning
confidence: 99%
“…Since the PAb used in the development of the immunoapproaches was able to recognise AZA carboxy cogeneres in addition to other AZA analogues, the raw mussel samples analysed were heated to perform LC-MS/MS analysis. Heating catalyses the decarboxylation of AZA carboxy congeners (e.g., AZA-17, AZA-19, AZA-21 and AZA-23) that may be present in the samples to AZA-3, AZA-6, AZA-4 and AZA-9 respectively [31,32] Figure 5), with no significant differences observed between the quantifications achieved by any of the immunoapproaches and LC-MS/MS analysis (P = 1.000).…”
Section: Azas Detection In Mussel Samplesmentioning
Given the widespread occurrence of azaspiracids (AZAs) it is clearly necessary to advance in simple and low-cost methods for the rapid detection of these marine toxins in order to protect seafood consumers. To address this need, electrochemical immunosensors for the detection of AZAs based on a competitive direct immunoassay using peroxidase-labelled AZA as a tracer were developed. An anti-AZA polyclonal antibody was immobilised in a controlled and stable manner on protein G or avidin-coated electrodes. Experimental conditions were first optimised using colorimetric immunoassays on microtitre plates, providing intermediate products already applicable to the accurate detection of AZAs. Then, transfer of the protein G and avidin-biotin interaction-based immunoassays to 8-electrode arrays provided compact and miniaturised devices for the high-throughput detection of AZAs. The low amounts of immunoreagents required as well as the potential for reusability of the avidin-biotin interaction-based immunosensors represented significant economic savings as well as a contribution to sustainability. The electrochemical immunosensors enabled the quantification of all regulated AZAs below the regulatory limit, as well as a broad range of other toxic AZA analogues (from 63 ± 3 to 2841 ± 247 µg AZA-1 equiv./kg for the protein G-based immunosensor and from 46 ± 2 to 3079 ± 358 µg AZA-1 equiv./kg for the avidin-biotin interaction-based immunosensor). The good agreement between the results obtained by the immunosensors and LC-MS/MS in the analysis of naturally contaminated mussel samples demonstrated the easy implementation of electrochemical immunosensors for routine analysis of AZAs in food safety monitoring programs.
“…Azaspiracids (AZAs) are nitrogen containing polyether toxins comprising a unique spiral ring assembly containing a heterocyclic amine (piperidine) and an aliphatic carboxylic acid moiety (FAO, 2005). Over 50 AZAs have been described in the literature (Rehmann et al., 2008; Kilcoyne et al., 2014, 2015a,b; Krock et al., 2015) of which some are found only in microalgae. Of all AZAs described in literature, AZA1 and AZA2 are the most widely found in shellfish, but beside these two also AZA3 is regulated (See Figure 4).…”
EFSA
was asked by the European Commission to provide information on levels of lipophilic shellfish toxins in whole scallops that would ensure levels in edible parts below the regulatory limits after shucking, i.e. removal of non‐edible parts. This should include the okadaic acid (
OA
), the azaspiracid (
AZA
) and the yessotoxin (
YTX
) groups, and five species of scallops. In addition,
EFSA
was asked to recommend the number of scallops in an analytical sample. To address these questions,
EFSA
received suitable data on the three toxin groups in two scallop species,
Aequipecten opercularis
and
Pecten maximus
, i.e. data on individual and pooled samples of edible and non‐edible parts from contamination incidents. The majority of the concentration levels were below limit of quantification (
LOQ
)/limit of detection (
LOD
), especially in adductor muscle but also in gonads. Shucking in most cases resulted in a strong decrease in the toxin levels. For
Pecten maximus
, statistical analysis showed that levels in whole scallops should not exceed 256 μg
OA
eq/kg or 217 μg
AZA
1 eq/kg to ensure that levels in gonads are below the regulatory limits of 160 μg
OA
or
AZA
1 eq/kg with 99% certainty. Such an analysis was not possible for yessotoxins or any toxin in
Aequipecten opercularis
and an assessment could only be based on upper bound levels. To ensure a 95% correct prediction on whether the level in scallops in an area or lot is correctly predicted to be compliant/non‐compliant, it was shown that 10 scallops per sample would be sufficient to predict with 95% certainty if levels of
OA
‐group toxins in the area/lot were 25% below or above the regulatory limit. However, to predict with a 95% certainty for levels between 140 and 180 μg
OA
eq/kg, a pooled sample of more than 30 scallops would have to be tested.
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