M astovska et al .: J ournal of aoaC I nternatIonal v ol . 98, n o . 2, 2015 503

qualification phase was the availability of resources and not the ability to qualify for the study.


Collaborators’ Comments

Most of the study participants commented very positively on the speed and ease of use of the method, especially laboratories currently analyzing PAHs with much more labor-intensive and time-consuming methods. The most frequently reported sources of PAH contamination were salts (which have to be muffled and stored appropriately). Some participants had problems with PAH sources in their laboratories caused by the use of oil pumps in the vicinity of the space used for the sample preparation. As noted above, one collaborator discovered PAHs in polypropylene centrifuge tubes used for practice sample analysis ( Note : All tubes and containers used for the test sample storage and preparation were pretested by the study direction team). Several collaborators initially had problems with optimization of the evaporation steps to prevent losses of volatile PAHs. As noted above, the addition of ethyl acetate in the second evaporation step resolved this issue in most cases. The majority of study participants used evaporation with a gentle stream of nitrogen at room temperature or a maximum of 40°C. Collaborators noticed differences in the color of extracts of oyster blank versus fortified samples stored for several months in a freezer at –20°C. The blank sample produced a dark green extract, whereas the same blank sample fortified with PAHs gave a yellow-brown extract. This was not observed for oyster samples stored for a shorter period of time and/or stored at –70°C. As discussed below, this observation could be linked to degradation issues in oysters. Also, the participants noted that oyster extracts were generally dirtier than shrimp and mussel extracts, which affected chromatography in some cases. One collaborator reported the use of a mechanical shaker instead of hand-shaking. Mechanical shaking is generally preferred by routine testing laboratories, and this method modification is acceptable as long as a vigorous and effective shaking (up and down in the tube) is ensured. Tables 3–11 provide the collaborative study results obtained by the participating laboratories in three blind duplicates (low, mid, and high fortification level) in shrimp, mussel, and oyster. Results from Laboratory No. 10 are presented only for mussel because the Study Directors excluded their oyster and shrimp data sets due to calibration (standard preparation) issues. 1,7-DMP was used as a homogenization check and was added to blank mussel and oyster samples at 40 and 80 µg/kg, respectively, during the homogenization step. The mean concentration value obtained for 1,7-DMP by all participants (except for sample SFC M4 lost by Laboratory No. 6 and the result for sample SFC M3 from Laboratory No. 1, which was removed as an apparent outlier) in all seven test mussel samples (three blind duplicates and one blank) was 38.8 µg/kg (RSD = 21.5%, n = 68), which corresponds to mean recovery of 97.0%. In the case of oysters, the mean concentration value obtained for 1,7-DMP by all participants (except for Laboratories 1 and 10, for which all 1,7-DMP results were eliminated as outliers in the Grubbs’ tests applied Collaborative Study Results



Figure 1. Structures of anthracene (Ant), benzo[ a ]anthracene (BaA), and benzo[ a ]pyrene (BaP).

Figure 1. Structures of anthracene (Ant), benzo[ a ]anthracene (BaA), and benzo[ a ]pyrene (BaP).

blank PAH levels were too high to participate in the collaborative study and to conduct low-level PAH analysis in general. This was typically due to their location (high environmental contamination) and/or their laboratory contamination. Some participants were able to reduce the reagent (procedure) blank contamination by moving the sample preparation (extraction, cleanup, and evaporation) away from oil pumps, such as MS rough pumps. The method requires heating of the used salts and recommends heating of glassware. Solvents, plastic material, and equipment may also be sources of PAHs, including polypropylene centrifugation/extraction tubes. As one collaborator discovered, simple testing of polypropylene tubes using ethyl acetate wash/extraction may not reveal PAH contamination. However, the extraction dynamic during the actual procedure can release potentially present PAHs into the extract when the tubes get heated due to the exothermic reaction caused by addition of MgSO 4 to the water-containing extraction mixture. For this and other potential contamination reasons, it is highly important to analyze a reagent blank with every sample batch. In addition to the contamination issues, another problem faced by laboratories less experienced in PAH analysis was optimization of the evaporation conditions to prevent losses of volatile analytes, especially naphthalene. Isooctane is used as a keeper in both evaporation steps, but it did not prevent significant losses of volatile PAHs in the second evaporation step in certain laboratories. For this reason, the study direction team recommended addition of 1–2 mL of ethyl acetate to the SPE eluent for a better control of the final evaporation process, which helped in most cases and was added as a recommendation to the method procedure. Sixteen laboratories entered the qualification phase, but only 10 of them (listed in the Acknowledgments section) completed the qualification successfully and/or continued in the study. In many cases, the reason why a participant did not complete the

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