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M

astovska

et al

.:

J

ournal of

aoaC I

nternatIonal

v

ol

.

98, n

o

. 2, 2015

503

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

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.

Collaborative Study Results

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

Figure 1. Structures of anthracene (Ant), benzo[

a

]anthracene (BaA), and

benzo[

a

]pyrene (BaP).

BaP

Ant

BaA

Figure 1. Structures of anthracene (Ant), benzo[

a

]anthracene

(BaA), and benzo[

a

]pyrene (BaP).