Techniques for Determination of Benzo(a)pyrene in Marine Organisms and Sediments Bruce P. Dunn' Cancer Research Center2, and.Department of Zoology, 2075 Wesbrook Place, University of British Columbia, Vancouver, B.C., Canada V6T 1W5
Rapid, economical, and reliable procedures are developed for the measurement of the carcinogen benzo(a)pyrene in small marine tissue and sediment samples. These involve alkaline digestion of samples, column chromatography on Florisil, DMSO extraction, separation of polycyclic aromatic hydrocarbons by thin-layer chromatography, and measurement of benzo(a)pyrene by fluorimetry. Recoveries of compound are measured for each sample by the use of radioactive benzo(a)pyrene as an internal standard. The procedures, which have a sensitivity of 0.1 yg/kg and a precision of 6%, appear more than adequate for application in routine monitoring programs for polycyclic aromatic hydrocarbon carcinogens in the marine'environment. The presence of polycyclic aromatic hydrocarbons (PAH) in the marine environment is well established. These compounds, many of which are strong carcinogens, have been found both in bottom sediments ( I , 2 ) and in organisms (3-6). They can enter the oceans by many routes, including petroleum spills, runoff from roads, sewage, effluents from industrial processes, and fallout from the atmosphere. It has been suggested that in addition, these compounds may also be biosynthesized (7,8), although there is evidence contrary to this view (9, 10). We have been investigating the use of the level of PAH in coastal waters as an index of the carcinogenic load on aquatic organisms. This topic has importance with respect to both the possibility of adverse effects on aquatic organisms living in contaminated waters and the public health aspects of contaminated seafoods. We have recently shown that the tissue level of the carcinogen benzo(a)pyrene (B(a)P) in mussels ( M y t h s edulis or M . californianus) may represent a good indicator for the contamination of bodies of water by PAH. There was a marked correlation between the presence of human activity in coastal areas and the contamination of mussels by B(a)P. Wharfs and docks and structures containing creosoted pilings or timbers appeared to be particularly strong sources of carcinogenic compounds (11). When creosote-contaminated mussels were transferred to clean water, they lost B(a)P exponentially, with a half-life of approximately two weeks (12). The problem of the origin, distribution, and fate of PAH in the marine environment has been complicated by the lack of methods of analysis that can be applied to the measurement of trace amounts of these compounds on a routine basis. This communication describes a rapid, sensitive, and reliable technique which can be used to measure the carcinogen benzo(a)pyrene in marine organisms and in bottom sediments. The technique is a small-scale, simplified version of the procedure described by Howard et al. for the measurement of B(a)P in foodstuffs ( 1 3 , 1 4 ) .A major feature of the modified procedure is the use of radioactively labeled benzo'(a)pyrene as an internal standard to enable the precise measurement of losses of compound during purification of each sample. 'Research Fellow of the National Cancer Institute of Canada. *To which correspondence should be addressed. 1018
Environmental Science & Technology
Reagents Solvents were practical or technical grade for economy and were redistilled before use in an all-glass still with a Vigreaux reflux column. Solvents were stored in glass-stoppered containers and were protected from contact with rubber or plastic a t all times (15). Their purity was checked by evaporating a suitable volume to 0.1 ml and examining the concentrated material for fluorescence under ultraviolet light. DMSO was spectral grade and was used without further purification, while hexadecane was practical grade and was purified before use by passage through a column of activated silica gel (16). Radioactively labeled benzo(a)pyrene (3,4-Benz(3,614C) pyrene, specific activity 21 mCi/mmol, or G-3H-Benzo(a) pyrene, specific activity 25 Ci/mmol) was purchased from Amersham/Searle and routinely purified before use by thinlayer chromatography on cellulose-acetate, Sodium sulfate and KOH were reagent grade and were used without further purification. Florisil (60-100 mesh) was from Matheson Coleman & Bell. Batches, 250 g, of the adsorbent were thoroughly washed by decantation with distilled water to remove fines and sodium sulfate (17).The wet material was placed in a 600-ml fritted glass Buchner funnel, excess water was removed by suction, and the adsorbent washed in the funnel by stirring with two 300-ml portions of methanol. Excess methanol was removed by suction, and the adsorbent dried overnight in vacuo at 60". The cleaned Florisil was activated at 250" for 18 h, then cooled and partially deactivated by the addition of 2% water. Cellulose-acetate (20% acetylated) for thin-layer chromatography was from Macherey-Nagel and Co. Thin-layer plates were formed by coating a 0.5-mm layer of cellulose-acetate (25 g in 100 ml of ethanol) on glass plates 10 X 20 cm. Plates were cleaned before use by development in the running solvent. Procedures Sample Preparation and Storage. Mussels 4-6 cm in size were collected from the midintertidal zone, cleaned externally, and then shucked. Byssal threads were removed, and excess fluid was drained away by agitating the tissue for 1min over a screen of mesh size 1.5 mm. Tissues from clams, oysters, and fish were prepared with appropriate modifications. Sediment samples were placed in clean glass containers and thoroughly mixed. A representative portion (4-5 g) was dried a t 80' for 48 h for the determination of moisture content. All biological specimens and sediments were stored at -10' until analysis. Extraction. To avoid possible photodecomposition of PAH (18),all extraction and purification procedures were carried out under subdued yellow tungsten light. Twenty to forty grams of tissues were placed in a 300-ml round-bottomed flask, and 150 ml of ethanol, 7 g of KOH, two or three boiling chips, and an aliquot of radioactive benzo(a)pyrene (either 1000 dpm 14C-B(a)P,ca. 5 ng, or.25 OOO dpm 3H-B(a)P,ca. 0.1 ng) were added. The tissue was digested by refluxing gently for 1.5 h with occasional swirling to prevent sticking to the bottom of the flask. The digest was added while hot to 150 ml of water in a 2-1. sep. funnel, and the digestion flask rinsed out with an additional 50 ml of ethanol. The water/ethanol mix was extracted 3 times with 200 ml of isooctane, and the iso-
octane extracts were combined and washed with 4 X 200 ml warm (60') water. Wet sediment samples weighing 10-20 g were refluxed in 100 ml of ethanol with 5 g of KOH, boiling chips, and radioactive tracer. The contents of the flask were then poured into a 250-ml Erlenmeyer flask, and the sediment was allowed to settle by gravity for 5 min. The supernatant was decanted through a glass wool plug into 150 ml of water in a sep. funnel. The sediment was washed by decantation with two additional portions of ethanol (50 ml each) which were also added to the sep. funnel. The remainder of the processing was as for tissue samples. Column Chromatography on Florisil. A column of 30 g of Florisil covered with 60 g of Na2S04 was prepared in a glass column (40 X 400 mm) with a coarse fritted glass disc. The column was prewashed with 100 ml of isooctane, and the isooctane extract of the sample then passed through. The column was allowed to drain and was then washed with two 100-ml portions of fresh isooctane, allowing the column to drain briefly between each addition. All isooctane passed through the column was saved for purification by distillation and eventual reuse. Polycyclic aromatic hydrocarbons were eluted from the column with 3 X 100 ml benzene. The combined eluate was reduced to 5 ml by rotary evaporation, 50 ml of isooctane was added, and the volume again reduced to 5 ml to remove the benzene. Safety note: Benzene is toxic to blood forming tissues. Procedures utilizing benzene should be carried out only with adequate ventilation. DMSO Extraction. PAH were extracted from the 5 ml of isooctane with 3 X 5 ml DMSO. The DMSO extracts were combined with 30 ml of water, and the PAH extracted into 2 X 10 ml isooctane. The isooctane extracts were combined, washed 3 times with 20 ml of water, and dried by passage through 10 g of Na2S04 in a 15-ml coarse fritted glass Buchner funnel. Thin-Layer Chromatography. The extract was reduced in volume to approximately 0.1 ml using rotary evaporation and then a stream of nitrogen and was applied as a narrow streak to the origin of a cellulose-acetate thin-layer plate. A standard of 10 ng B(a)P was applied as a spot to one side of the plate, and the plate was developed with ethanol/toluene/water (17/4/4) (16). The plate was positioned in the development tank with the bottom of the plate supported approximately 5 cm from the base of the tank on a row of beakers inside the tank. Sufficient developing solvent was present to cover the beakers and wet the bottom of the plate. The lid of the tank was left slightly ajar, with the top of the plate extending approximately 2 cm through the gap into the open air. Development was continued for 5-6 h. The solvent front reached the top of the plate in about 2 h, and subsequent development occurred due to evaporation of the developing solvent at the exposed top of the plate. At the end of the development, the plates were partially air-dried, and the B(a)P band was located and outlined under long-wave ultraviolet light. The B(a)P band, with an Rf of approximately 0.3 after 2 h development, was always the lowest fluorescent band on the plate. The adsorbent at the position of the B(a)P band was scraped off the plate while still damp and placed in a 15-ml fine fritted glass Buchner funnel. The B(a)P was removed from the cellulose-acetate by washing with 4 X 4 ml hot (65") methanol, using gentle suction. The methanol was added to 10 ml of a solution of 20% hexadecane in isooctane, and the methanol and isooctane were removed by rotary evaporation to leave the B(a)P in 2 ml of hexadecane, ready for fluorimetry. Fluorimetric Measurement of B(a)P. B(a)P was measured fluorimetrically in hexadecane using the baseline
technique of Kunte (19). Saqples and standards of 10-200 ng B(a)P/ml in hexadecane were excited a t 365 nm in an Aminco-Bowman spectrophotofluorimeter, and the emission spectrum was recorded from 375 to 500 nm. An artificial baseline was drawn between minima in the fluorescence spectrum occurring at 418 and 448 nm, and the height of the peak at 430 nm above this baseline was measured. When necessary, highly fluorescent samples were diluted with hexadecane to bring their fluorescence within the range of the standards used. Calculation of Percentage Recovery and Amount of B(a)P Originally Present in Sample. After fluorimetry, the amount of radioactive B(a)P internal standard in each sample was determined by scintillation counting. The recovery of B(a)P was calculated by comparing the amount of radioactivity added at the beginning of the digestion procedure with the amount recovered in the fluorimetry sample. The amount of B(a)P determined by fluorimetry was then corrected if necessary for the contribution of radioactive tracer (this correction is negligible if the higher specific activity 3H-B(a)P is used), and the net amount of B(a)P was then corrected for losses in the procedures. The amount of B(a)P originally present in the sample was then expressed as pg B(a)P/kg wet weight of tissue or dry weight of sediment.
Results Notes on Procedures. The overall recovery of B(a)P was generally 60-80% for tissue samples (mussels, clams, oysters, or fish) and 50-70% for sediment samples. Recoveries were the same whether 14Cor 3H labeled B(a)P was used as an internal standard. Recoveries of B(a)P from the alcoholic KOH digest plus water into isooctane were essentially quantitative for tissue samples and generally over 90% for sediment samples. With the proportions of ethanol and water used, emulsion problems at this step were generally negligible. Occasional slow separations could be speeded by the addition of 10-20 ml of a saturated solution of NaCl in water. Recoveries from the Florisil column were generally over 90%, but fluctuated somewhat depending on the nature of the samples and the method of preparation of the Florisil. Losses occurred by tailing of B(a)P on the column during elution with benzene, rather than by failure of B(a)P to be retained by the column during elution with isooctane. The problem of tailing was accentuated with unusually small samples, samples with only a low extractable organic content (such as some sandy sediments), or samples containing only very small amounts of B(a)P. Florisil pretreated as described by Howard et al. (16) gave somewhat irreproducible results. In an attempt to better standardize various batches of Florisil, the adsorbent was first extensively washed with water to remove sodium sulfate, a contaminent of commercial Florisil which can cause varying adsorbent activity (17). The adsorbent was then washed with methanol to remove organic impurities, dried, and activated. Fully activated Florisil was unsuitable as a chromatography substrate as it gave extensive tailing and poor recoveries, but the addition of 2% water to the activated material gave an adsorbent with consistently good chromatography properties. With the marine samples processed in this laboratory, the phosphoric acid extraction step described by Howard et al. (13)was unnecessary. In many cases the procedure could be even further simplified by eliminating the DMSO extraction step. Some tissue samples and most sediment samples, however, contained interfering materials not eliminated by column chromatography and which remained a t or near the origin of thin-layer chromatograms, causing extensive streaking of the migrating PAH. With these samples the interfering materials could be eliminated by a small-scale, simplified version of the DMSO extraction procedure described by Howard et al. (13). Volume 10, Number 10, October 1976
1019
Because there was no certain way to identify those samples requiring DMSO extraction, this procedure was eventually adopted for all samples. Dimethyl formamide (20) was unsuitable as a substitute for DMSO since extraction procedures utilizing it failed to remove interfering materials from extracts. With all samples the violet fluorescing B(a)P band on thin-layer plates was associated with a blue fluorescing band with a slightly greater Rj value. This band was only partially resolved from the B(a)P band during normal chromatography but was completely resolved from the B(a)P by using extended chromatography (methods).The Rj value of this band, as well as the ultraviolet and fluorescence spectra of material isolated from this band, was identical to those of an authentic sample of benzo( b)fluoranthene. If the total fluorescence at a fixed wavelength is used as a measure of B(a)P concentration, the determination is subject to errors in the presence of any background fluorescence. For this reason, B(a)P was measured by using the baseline technique of Kunte (19), which involves determining the height of one of the peaks of the B(a)P emission spectrum above an artificial baseline drawn between two minima in the B(a)P spectrum. This technique is analogous to baseline techniques widely used for the determination of PAH by their ultraviolet absorption. Reproducibility of B(a)PDetermination. The precision of the analytical technique was examined by analyzing replicate samples of mussel tissue, prepared from mussels from three geographical locations known to be contaminated with B(a)P. For each site the total tissue from 200 to 300 mussels was homogenized in one container. Subsamples of 20-30-g tissue were prepared from each homogenate and either analyzed immediately or stored at -10’. The standard deviation of the analytical results ranged from 3.2 to 8.1%of the mean, with a n average of 6.2% (Table I). Samples stored for 12 weeks a t -10’ showed no statistically significant change in B(a)P content, suggesting that this method of storage is suitable for periods of at least several months. The reproducibility of B(a)P level in field samples of mussels was examined by analyzing six separate samples (total tissue from 10-15 mussels each) from each of several locations. Variability in this case reflects not only variability in the measurement technique itself, but also differences in the actual content of B(a)P in individual field samples. Table I1 shows that the level of contamination of naturally occurring mussels can vary widely. The higher levels in the last three groups of samples probably reflect direct contamination from creosote (11).At both high and low levels, standard deviations approximated 20% of the mean B(a)P level (average 20.8%, range 9.2-31.4%.
Discussion Considerable tailing of B(a)P occurred on highly activated Florisil columns and when samples were unusually small. Snyder (21) reported that fully activated Florisil contains a proportion of highly active acidic sites which can chemisorb PAH such as perylene, causing tailing on columns. He found that these sites could be selectively blocked by the addition of a t least 1%water to the adsorbent. Similar results were obtained in the current study. Fully activated Florisil3retained substantial amounts of B(a)P during elution with benzene, whereas the same adsorbent deactivated with 2% water gave nearly quantitative recoveries. That samples with a low organic content frequently have somewhat lower recoveries of B(a)P from the Florisil column suggests that organic compounds present in sample extracts may similarly block the active sites responsible for the tailing of B(a)P. The use of a radioactive internal standard with each sample processed lends considerable reliability to the analytical 1020
Environmental Science & Technology
Table 1. Precision of Assay for Benzo(a)pyrene B(a)P level, pg/kg wet wt Area
Styrofoam floats, marina “A” Untreated wood floats, marina
na
SD as %
19.9 1.35 1 1.9 0.38
6 4
6.8 3.2
18.4 1.24
6
6.7
17.8 1.45
4
8.1
Mean
6.2
Mean
SD
“0”
Concrete, 2-4 m from group qf 30 creosoted pilings As above, stored 12 weeks at
-loo a
Subsamples of a homogenate of tissue from 200 to 300 mussels from each
location.
Table II. Reproducibility of Benzo(a)pyrene Level in Field Samples of Mussels B(a)P level, wglkg wet weight Area
Mean
SD
na
Shoreline, outer Vancouver harbor 0.55 0.11 6 200 m from a moderate-sized 1.78 0.40 6 marina On group of 30 creosoted pilings 49.2 15.4 6 On concrete, 2-4 m from above 45.3 4.2 6 location As above, 2 weeks later 49.1 10.4 6 Mean a
SDas %
20.0 22.3 31.4 9.2 21.1 20.8
Individual samples comprised of tissue from 10 to 15 mussels each.
procedure. Although the use of an internal standard has many advantages and takes little additional operator time, it has to date been used only infrequently for the analysis of PAH (22-24). When using a radioactive tracer of a fixed known specific activity, the results of the analysis for B(a)P depend on only three parameters: the amount of radioactive tracer added, the amount of tracer recovered (determined by scintillation counting), and the total amount of B(a)P isolated from the sample (determined fluorimetrically). It is important to emphasize that changes in procedure (deliberate or otherwise) which affect only the recovery of the B(a)P through the procedure do not affect the end result. This gives a very “robust’’ technique which is resistant to errors, changes in reagents, and variations in technique between different operators. An additional advantage is the elimination of the timeconsuming procedure of performing separate recovery studies each time procedures are altered or a new sample type is processed. Note that neither recovery studies involving the spiking of uncontaminated samples with known amounts of B(a)P (13, 25) nor the use of a radioactive internal standard added to the sample digestion flask can correct for any incompleteness in the extraction into solution of tissue or particle associated B(a)P. The efficiency of extraction can only be estimated by comparison with other extraction procedures. Farrington and Medeiros have reported (27)that digestion of shellfish tissue in refluxing KOH and methanol for 1 h yields slightly less hydrocarbons than a 48-h Soxhlet extraction with methanokbenzene. For a routine method, however, alkaline digestion is much more suitable than Soxhlet extraction since it is faster, simpler, does not require special glassware, and does not require subsequent separate saponification of sample lipids. When using high specific activity SH-B(a)Pas an internal standard where the amount of tracer is negligible in comparison with the amount of B(a)P from the sample, the sensitivity of the technique is limited only by the sensitivity of the fluorimetric measurement. Concentrations of B(a)P in hexadec-
ane on the order of 1ng/ml were easily measured in a standard Aminco-Bowman fluorimeter with 1-cm2 cells, and lower concentrations could undoubtedly be measured using more sophisticated instrumentation. Assuming 80% recovery, a 2-ml fluorimetry sample, and a 25-g sample, this corresponds to a sample contamination of 0.1 pg/kg. Higher sensitivities could be obtained by increasing the sensitivity of the fluorimetric measurement, increasing the size of the sample, or both. The sample extraction and purification procedures reported here are directly based on the pioneering work of Howard et al. (13, 14, 16) on the measurement of polycyclic aromatic hydrocarbons in foodstuffs. The original procedures have been modified in the interest of speed, simplicity, reliability, and economical use of reagents. Howard and coworkers, reporting on a collaborative study in which a number of different laboratories measured the level of B(a)P in food samples spiked with 0.4 or 1.0 pg of the compound, reported a standard deviation for fluorescence measurements of between 3.24 and 10.8% within individual laboratories and between 5.18 and 13.6%when comparing different laboratories (25).Malanoski et al., using the same technique, found a standard deviation of 25% for a series of meat samples spiked with 0.2 kg B(a)P (26). In comparison, the variability found in the current study ranged from 3.2 to 8.1%comparing favorably with data from the original technique. An analysis of sources of variability in the current study suggests that the major reason for lack of precision was nonreproducibility of fluorimetric measurements of B(a)P. The analytical procedure for benzo(a)pyrene takes 3-4 h of operator time per sample, including time taken for the preparation of reagents. Reagent costs are estimated to be less than one-half that of the procedure described by Howard et al. (13,14). The technique has more than enough sensitivity to measure B(a)P in environmental samples (11).The results of the analysis of field samples suggest that the precision of the measurement of B(a)P is 3-4 times better than the variability inherent in the samples being processed. This, coupled with the fact that the level of B(a)P in mussels taken from various areas may vary by more than two orders of magnitude ( I I ) , suggests that the method is entirely adequate for use in a routine monitoring program for PAH in the marine environment. Acknowledgment
I thank H. F. Stich and C. T. Beer for advice during the work and the preparation of the manuscript, D. Hoffmann and
E. Sawicki for the generous gift of polycyclic aromatic hydrocarbon reference compounds, and W. Walsh for expert technical assistance.
Literature Cited (1) Giger, W., Blumer, M., Anal. Chem., 46,1663-71 (1974). (2) Bhmer, M., Youngblood, W. W., Science, 188,53-55 (1975). (3) Cahnmann, H. J., Kuratsune, M., Anal. Chem., 29, 1312-17 (1957). (4) Zechmeister, L., Koe, B. K., Arch. Biochem. Biophys., 35,l-11 (1952). (5) Suess, M. J., Arch. Hyg. Bakteriol., 154,l-7 (1970). (6) ZoBell, C. E., Proceedings of the Joint Conference on Prevention and Control of Oil Saills. DD 441-51. American Petroleum Institute Washington, D.C., i971: ( 7 ) Borneff, J., Selenka, F., Kunte, H., Maximos, A., Enuiron. Res., 2,22-29 (1968). (8) Graf, W., Diehl, H., Arch. Hyg. Bakteriol., 150,49-59 (1966). (9) Youngblood, W. W., Blumer, M., Geochim. Cosmochim. Acta, 39,1303-14 (1975). (10) Grimmer, G., Duvel, D., Z. Naturforsch., 25b, 1171-75 (1970). Exp. Biol. Med., 150,49-51 (11) Dunn, B. P., Stich, H. F., Proc. SOC. (1975). (12) Dunn, B. P., Stich, H. F., Bull. Enuiron. Contam. Toxicol., 15 398-401 (1976). (13) Howard, J. W., White, R. H., Fry, B. E., Turicchi, E. W., J. Assoc. Off. Anal. Chem., 49,611-17 (1966). (14) Association of Official Analvtical Chemists. J. Assoc. Off. Anal. ' Chem., 51,449-53 (1968). (15) Kordan, H. A., Science, 149,1382-83 (1965). (16) Howard, J. W., Teague, R. T., White, R. H., Fry, B. E., J. Assoc. Off. Anal. Chem., 49,595-611 (1966). (17) Mills, P. A., ibid., 51,29-32 (1968). (18) Kuratsune, M., Hirohata, T., Nut. Cancer Inst. Monogr., 9, 117-25 (1962). (19) Kunte, H., Arch. Hyg. Bakteriol., 151, 193-201 (1967). (20) Smith, C. G., Nau, C. A., Lawrence, C. H., Am. Ind. Hyg. Assoc. J., 29,242-47 (1968). (21) Snyder, L. R., J . Chromatogr., 12,488-509 (1963). (22) Hoffmann, D., Wynder, E. L., Cancer, 15,93-102 (1962). (23) Seaman, W., Anal. Chem., 29,1570-73 (1957). (24) Pancirov, R. J., Brown, R. A., Proceedings of the 1975 Conference on Prevention and Control of Oil Pollution, pp 103-13, American Petroleum Institute, Washington, D.C., 1975. (25) Howard, J. W., Fazio, T., White, R. H., J . Assoc. Off. Anal. Chem., 51,544-48 (1968). (26) Malanoski, A. J., Greenfield, E. L., Barnes, C. J.,Worthington, J. M., Joe, F. L., ibid., pp 114-21. (27) Farrington, J. W., Medeiros, G. C., Proceedings of the 1975 Conference on Prevention and Control of Oil Pollution, pp 115-21, American Petroleum Institute, Washington, D.C., 1975.
Received for review January 6, ,1976.Accepted April 20, 1976.
Volume
IO, Number 10, October 1976
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