ANALYTICAL CHEMISTRY, VOL. 50, NO. 6, MAY 1978
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Determination of Trace Level Hydrocarbons in Marine Biota S. N. Cheder," B. H. Gump, H. S. Hertz, W. E. May, and S. A. Wise Institute for Materials Research, Analytical Chemistry Division, National Bureau of Standards, Washington, D.C. 20234
A method is described for the determination of petroleum hydrocarbons in marine biota. This procedure utilizes dynamic headspace sampling of an aqueous caustic tissue homogenate to extract and collect volatile organic components. Interfering polar biogenic (non-anthropogenic) components are removed by normal-phase high-performance liquid chromatography. Quantitation and identification of the individual compounds are accomplished using gas chromatography and gas chromatography-mass spectrometry. The nonvolatile polynuclear aromatic hydrocarbons which remain in the homogenate afler headspace sampling are solvent-extracted and then analyzed by reversed-phase liquid chromatography.
The low concentration of hydrocarbons in marine petroleum pollution baseline and monitoring studies necessitates the development of analytical techniques for biota which are sensitive a t the Fg/ kg level. Accurate determination of these low hydrocarbon levels requires t h e solving of significant problems. Since most of the organic compounds present are of biological origin, a suitable chemical cleanup of the sample is necessary in order t o remove interfering nonhydrocarbon compounds prior to t h e measurement of t h e low levels of hydrocarbons present. In addition, i t is important to know the extraction efficiency of the petroleum hydrocarbons from the tissue sample so that appropriate sample sizes can be used for t h e desired sensitivity levels. Furthermore, analytical methods which ultimately permit the identification of individual hydrocarbon components are desired. Presently, a variety of analytical techniques exist for the determination of hydrocarbons in marine tissue (1-9). T o remove t h e hydrocarbons from t h e tissue matrix, these methods generally employ Soxhlet extraction (1-3),alcoholic or aqueous caustic digestion followed by organic solvent extraction ( 4 @ , or homogenization/sonication in the presence of an organic solvent (7). Farrington and Medeiros (8)recently compared the efficiencies of these various extraction procedures and found them to be approximately equal for hydrocarbon extraction. In all of the above procedures, t h e hydrocarbons extracted from the tissue matrix are generally isolated by silica gel-alumina column chromatography and quantified by gravimetric or gas chromatographic analysis. A different procedure utilizing gas chromatographic analysis of the headspace vapor collected over a heated tissue sample has been employed by Deshimaru (9) t o monitor t h e hydrocarbons in fish exposed t o crude oil. A rapid and sensitive method for the determination of hydrocarbons in marine biota has been developed in this laboratory. This method is schematically represented in Figure 1. The method utilizes dynamic headspace sampling of an aqueous caustic tissue homogenate to extract and collect the volatile organic components. Interfering polar biogenic (non-anthropogenic) components are removed by high-performance liquid chromatography (HPLC) prior to quantitation and identification of t h e petroleum hydrocarbons by gas chromatography (GC) and gas chromatography-mass spectrometry (GC/MS). The nonvolatile hydrocarbon components e.g., PAHs) are solvent-extracted from the caustic tissue This paper not subject to U S . Copyright
homogenate after headspace sampling and then analyzed by reversed-phase HPLC with ultraviolet (UV) and fluorescence detection. This procedure provides the sensitivity necessary to determine low levels of hydrocarbons in marine organisms from relatively pristine environments.
EXPERIMENTAL Preparation of Marine Tissue Samples. Because of the low levels of hydrocarbons present in the samples, special caution was exercised to avoid contamination during collection and handling of the tissue samples (IO). All samples were quick-frozen in the field and were stored in a freezer at -10 "C until analyzed. All analyses to date have been performed on marine bivalves. At the time of analysis, samples were allowed to thaw only enough to facilitate removal of the tissue from the shell without loss of the surrounding fluid. Samples were prepared for analysis in a laminar flow hood in a cold room (4 "C). The procedure developed in this laboratory for the determination of hydrocarbons in marine tissue utilizes a modified version of the dynamic headspace sampling technique previously described for the analysis of marine sediments and water ( 1 0 , I I ) . Approximately 30 g of tissue, 500 mL of hydrocarbon-free water, and 50 g of NaOH were combined in a 2-L, 3-neck, round-bottom flask. An internal standard solution of aromatic and/or aliphatic hydrocarbons (-2 pg each) was added to the mixture and the contents of the flask were homogenized for -1.5 min using an ultrasonic probe (Brinkmann Polytron PT 10 20 340D) inserted in the center neck of the flask. The tissue homogenate was heated to 70 "C and headspace sampled for -18 h at a nitrogen flow of approximately 150 mL/min. The headspace vapors were passed through a 6.5 X 0.6 cm 0.d. stainless steel column packed with approximately 0.2 g TENAX-GC (Applied Science Laboratories, State College, Pa.). For headspace sampling, nitrogen was blown across the liquid interface rather than bubbled through the solution to avoid frothing of the flask contents. Following the sampling period, the TENAX-GC column was connected directly to a dry nitrogen stream (flow rate = -150 mL/min) for 4 h to remove water from the column. The column was then capped tightly and stored at 4 "C for subsequent HPLC cleanup. The homogenate solution remaining in the flask after headspace sampling was extracted with -30 mL of freshly distilled pentane to remove the nonvolatile PAHs. Chromatographic Procedures. Following headspace sampling and drying, the TENAX-GC column was connected as part of the injection loop of a liquid chromatograph, and the organic compounds were eluted from the TENAX-GC column with freshly distilled pentane onto a 30 cm X 4 mm i.d. pBondapak NH, column (Waters Associates, Milford, Mass.). The first 15-mL fraction from the pBondapak NH2 column contained the hydrocarbons. This hydrocarbon fraction was collected in a clean 15-mL centrifuge tube and then reduced to -300 pL by passing a stream of nitrogen over the solution. The nitrogen was purified by passing it through a trap containing molecular sieves. The concentrate was then transferred onto a clean TENAX-GC column using a 100-pL syringe. The centrifuge tube was rinsed twice with -50 pL of pentane and the washings were also transferred onto the column. A blank, which consisted of 15 mL of pentane concentrated by passing a stream of nitrogen over it, resulted in no detectable impurities, indicating no contamination from either the solvent or the nitrogen gas. The gas chromatographic system and procedure used for the determination of the hydrocarbons trapped on the TENAX-GC column have been described previously ( 1 0 , I I j . This procedure was modified slightly to facilitate the removal of the excess Published 1976 by the American Chemical Society
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pentane by allowing a flow of -35 mL/min of helium to pass through the TENAX-GC column for 10 min, before connecting it to the glass SE-30 coated, SCOT analytical column. GC analyses were performed with a 4-min isothermal period at 80 "C followed by temperature programming at a rate of 4 "C/min to 275 "C and h o1ding. The 30-mL pentane extract obtained by extracting the headspace-sampled tissue homogenate was concentrated to 200 p L by blowing a stream of nitrogen over the solution. Approximately 1 mL of acetonitrile was then added and the pentane removed by again passing nitrogen over the solution. The sample was then analyzed on a reversed-phase pBondapak CI8 liquid chromatographic column (30 cm X 4 mm i.d., Waters Associates, Milford, Mass.) utilizing both UV absorption and fluorescence emission detection. The fluorescence emission detector was an Aminco Filter Fluorometer (American Instrument Co., Silver Spring, Md. 20910) with an excitation filter with maximum transmission at 365 nm and an emission cut-off filter that passes wavelengths above 415 nm. A linear gradient elution profile from 50-100% acetonitrile in water at 2%/min was used for the analysis.
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RESULTS AND DISCUSSION One area of concern in t h e development of an analytical method for marine biota was the relative recoveries of the petroleum hydrocarbons present in a tissue sample. T h e addition of an internal standard a t the beginning of the sample workup enables one t o correct for losses which may occur during the analytical scheme. (This is in direct contrast to other procedures ( 3 , 6 )which add the internal standard just prior to the final concentration and chromatographic analysis.) Aromatic and aliphatic hydrocarbons investigated for use as internal standards in this study were mesitylene, naphthalene, 2,3,6-trimethylnaphthalene, phenanthrene, 2-methylundecane (Me(&), 5-methyltetradecane (Me(&), 7-methylhexadecane (MeC16),and 2-methyloctadecane (MeCI8). By using a mixture of compounds with a wide range of volatilities, it is possible t o correct for the varying recoveries over the range of molecular weights analyzed. Recovery data for t h e aromatic and aliphatic compounds used as internal standards in the tissue analyses are given in Table I. By extending the headspace sampling period from 4 h as previously reported for water and sediment (10, 11) to -18 h a t 70 "C, recoveries from water for the higher molecular weight aromatic and aliphatic components (Le., trimethyl-
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ANALYTICAL CHEMISTRY, VOL. 50, NO. 6, MAY 1978
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Table 11. Comparison of Volatile Hydrocarbon Levels Obtained with and without HPLC Cleanup, Mg/kg N o HPLC HPLC M y tilus (mussels)
1406 f 98' (2)d (Northeastern Gulf of Alaska) 540 ?- 46 ( 3 ) 1834 (1) Oysters (Middle Marsh, S.C.) ' 652 (1) Clams A (control) 509 i 11 ( 2 ) 377 t 88 (2) 491 t 8 8 ( 3 ) Clams B (1p g crude oil/g water)a 1421 t 114 ( 2 ) 1413 ?: 398 ( 2 ) Clams C ( 1 0 pg crude oil/g water)b 1704 (1) Sediment (collected near natural oil seep) 566 i 37 ( 2 ) 574 (1) Data reported as the standard deviaExposed to 10 Hg crude oil/g of water. a Exposed to 1 Mg crude oil/g of water, Denotes the number of samples analyzed. tion (1 U ) of a set of replicate values from the mean of the replicate values.
naphthalene, phenanthrene, MeCI6, and MeCI8) were increased to nearly 100%. The extended headspace sampling period also resulted in more reproducible recoveries. Hachenberg and Schmidt (12)have suggested the addition of an electrolyte t o increase the sensitivity of headspace analysis. However, the addition of potassium chloride to the tissue homogenate was found to have little effect on the internal standard recoveries. The addition of sodium or potassium hydroxide to digest the tissue matrix improved the aromatic hydrocarbon recoveries somewhat, but had no appreciable effect on the aliphatic hydrocarbon recoveries. Aliphatic hydrocarbon recoveries were found to be much lower than aromatic hydrocarbon recoveries in the headspace sampling of the tissue homogenate. Using the extended headspace sampling period and caustic digestion, recoveries from mussel tissue homogenate approached 100% for the higher aromatics but were only 30% for the aliphatic components. It is assumed that the aliphatic hydrocarbons are being retained in the lipophilic portion of the tissue homogenate and that the partition coefficient for these hydrocarbons between the headspace sampling gas and the lipophilic fraction is quite unfavorable. Recovery data for the complete analytical scheme indicate that some losses of the internal standards also occur during the liquid chromatographic cleanup and concentration step. The losses that occur during the concentration step amount to -25% for mesitylene, -30% for 2-methylundecane, -40% for naphthalene, 11% for 5-methyltetradecane, 5% for trimethylnaphthalene, and less than 1% for 7-methylhexadecane, 2-methylnaphthalene, phenanthrene, and hydrocarbons of higher molecular weight. Since quantitation in these analyses was dependent upon an internal standard added a t the beginning of the analytical scheme, it was imperative to know whether the internal standard components were recovered to the same extent as these components would be if incorporated in the tissue matrix. In a series of experiments with live Mytilus (mussels) exposed to 14C-naphthaleneand then analyzed using the 4-h headspace sampling procedure and no HPLC cleanup, a 14C recovery of 78 f 1 2 % was observed. In comparison, the recovery of nonlabeled naphthalene added as an internal standard was found to be 66 i 8% for the same 4-h headspace sampling procedure (see Table I). Indications are, therefore, that a t least in the case of naphthalene, an internal standard added to the mussel tissue solution can be recovered essentially to the same extent as naphthalene incorporated into live mussels. Preliminary GC/MS investigations of headspace sampled Mytilus homogenate indicated that the major biogenic compounds present on the TENAX-GC trap after headspace sampling were long chain aliphatic alcohols. Since these biogenic compounds interfere with the GC analysis of the petroleum hydrocarbons, several HPLC packing materials were investigated for their ability to separate petroleum hydrocarbons from the more polar biogenic compounds. The most satisfactory results were obtained using pBondapak NH2
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which has a highly polar aminosilane functional group chemically bonded to the support. This material is generally used for normal-phase HPLC to separate strongly polar compounds. However, when using a nonpolar mobile phase such as pentane, the pBondapak NH2 column provides a class separation similar to that obtained using a silica column, Le., saturated hydrocarbons elute before unsaturated hydrocarbons and aromatics, and the elution volume for aromatics increases with the number of condensed rings (13). Long chain alcohols are strongly retained on the pBondapak NH, column when pentane is used as the mobile phase and they can be removed from the column only after increasing the percentage of a more polar solvent (e.g., methylene chloride or methanol) in the mobile phase. The use of pBondapak NH2 eliminates a major problem encountered when using silica or alumina, Le., deactivation of the adsorbent by traces of water in the sample or mobile phase (resulting in a loss of resolution and nonreproducible separations). The dual trace (two different attenuations) gas chromatograms presented in Figure 2 for the analysis of a Mytilus sample illustrate the removal of the more polar biogenic components by this HPLC technique. Whereas the majority of the peaks are off-scale in the upper trace of Figure 2A, the intensities of the peaks in the upper trace of Figure 2B have been reduced considerably by HPLC cleanup. The numbered peaks in the chromatograms are the internal standards which were added to facilitate quantitation of the hydrocarbons. Quantitation of the hydrocarbons was accomplished using several internal standards as previously described (11). Figure 2C is the dual trace gas chromatogram of those components removed from the sample by the HPLC cleanup and subsequently eluted from the pBondapak NH, column with methanol. The effective removal of the more polar biogenic components by HPLC using a pBondapak NH, column is demonstrated in Table 11. These data from various tissue samples (mussels and clams) and a sediment sample indicate that HPLC removal of the nonhydrocarbon components is necessary to effectively determine low hydrocarbon levels in tissue. Of particular interest in Table I1 are the results obtained with various clam samples with and without exposure to 1 and 10 pg of crude oil/g of water. Dual trace gas chromatograms of these clam samples after HPLC cleanup are shown in Figure 3 (A, B, and C). A comparison of the data for the control clams with and without HPLC cleanup (Figures 3C and 3D, respectively) reveals that the six most abundant components (-100 pg/kg total) in the sample without HPLC cleanup are nonhydrocarbon. These six peaks are labeled a through f in Figure 3D. A comparison of the results obtained, after HPLC and after excluding the control level (Le., 400 pg/kg), for the clams exposed to 1 pg crude oil/g of water shows a difference of a factor of 10 in petroleum uptake. The data in Table I1 support the applicability of the above method for the determination of hydrocarbons in marine organisms exposed to toxic levels as well as those from unpolluted environments.
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ANALYTICAL CHEMISTRY, VOL. 50, NO. 6, MAY 1978
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TIME, min Figure 2. Dual trace (two different attenuations) gas chromatograms of headspace sampled Mytilus (A) without HPLC cleanup and (B) with HPLC cleanup. (C) Biogenic components removed from pBondapak NH, column with methanol after HPLC cleanup of Mytilus sample. Column: glass SE-30 coated SCOT column (100 m X 0.75 mm i.d.). Conditions: Helium at 6 mL/min, 4 min isothermal at 80 OC, then 4 OC/min at 275 OC and hold. Internal standards added for quantitation: (1) 2-methylundecane, (2) naphthalene, (3) 5-methyltetradecane, (4) 2,3,6-trimethylnaphthalene, (5) 7-methylhexadecane, (6) phenanthrene, (7) 2-methyloctadecane
T o evaluate the effect of the HPLC cleanup on petroleum hydrocarbons, a sediment sample with a natural low-level petroleum hydrocarbon burden and with essentially no biogenic contamination was analyzed with and without HPLC cleanup. The data in Tahle I1 for this sediment indicate that the measured petroleum hydrocarbon level determined by the headspace sampling procedure is, as expected, unaffected by the HPLC cleanup.
After removal of the volatile hydrocarbons by headspace sampling, the nonvolatile PAHs are determined by solvent extraction and HPLC/fluorescence analysis. The previously reported coupled-column technique for sediment and water ( 1 0 , I I ) was not employed here because of the incompatibility of t h e highly basic solution remaining after headspace sampling and the chemically-bonded reversed-phase packing material. A liquid chromatogram of the PAHs extracted from
ANALYTICAL CHEMISTRY, VOL. 50, NO. 6, MAY 1978
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Figure 3. Dual trace (two different attenuations) gas chromatograms of headspace sampled clams (after HPLC cleanup) exposed to crude oil at (A) 10 p g oil/g water, and (B) 1 p g oil/g water, and dual trace gas chromatograms of headspace sampled control clams, (C) with HPLC cleanup and (D) without HPLC cleanup. Column and Conditions: Same as in Figure 2, Internal standards added for quantitation: same as in Figure 2. Peaks a through f : nonhydrocarbon compounds removed by HPLC cleanup
a headspace-sampled solution of contaminated Mytilus tissue collected near Coal Oil Point, adjacent to natural oil seeps near Santa B a r b a r a , Calif., is shown in Figure 4. Individual PAHs can be identified by using normal-phase HPLC to isolate the
PAHs according to the number of condensed aromatic rings, and then subsequent analysis of the fractions by reversedphase HPLC with fluorescence emission detection as described by Wise et al. (13).
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ANALYTICAL CHEMISTRY, VOL. 50, NO. 6, MAY 1978
In contrast to previously reported methods for tissue, this headspace sampling procedure is applicable to the determination of low levels of hydrocarbons in relatively pristine areas as well as greater levels found in organisms exposed to petroleum. This procedure has been employed in this laboratory for numerous samples and found to be rapid and convenient for hydrocarbon determinations.
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Figure 4. Reversed-phase HPLC analysis of a solvent extract of the Mytilus homogenate after headspace sampling with (A) UV absorption detection at 254 nm and ( 8 )fluorescence emission detection above 415 nm
T h e headspace sampling procedure for the analysis of hydrocarbons in marine biota offers several advantages over current solvent extraction procedures. The headspace sampling technique requires minimal sample handling, few sample transfers, and only a minimal amount of organic solvent, thereby reducing the risk of contamination (a system blank for the headspace sampling method results in a value of only - 5 wg/kg based on a sample of 600 m L of water (10, 11)). In addition, only one solvent concentration step is involved, thereby reducing the losses of the more volatile components. When compared to solvent extraction procedures, the analyst's time is greatly reduced using the headspace sampling technique. During the lengthy headspace sampling period, the system is left to run unattended.
(1) M. Biumer, G. Souza, and J. Sass, Mar. Biol., 5 , 195 (1970). (2) R. F. Lee, R. Sauerheber, and A. A. Benson. Science, 177, 344 (1972). (3) R. C. Chrk, Jr., and J. S.Finley, "Proceedings of the 1973 Joint Conference on Prevention and Control of Oil Spills", American Petroleum Institute, Washington, D.C., 1973, p 161. (4) J. W. Blaylock, P.W. O'Keefe, J. N. Roehm, and R. E. Wilding, ibid. p 173. (5) R. J. Pancirov and R. A. Brown, "Proceedings of the 1975 Conference on Prevention and Control of Oil Pollution", American Petroleum Institute, Washington, D.C., 1975, p 103. (6) J. S. Warner, Anal. Chem., 48, 578 (1976). (7) J. J. Stegeman and J. M. Teal, Mar. Bioi., 22, 37 (1973). (8) J. W. Farringon and G. C. Medeiros, "Proceedings of the 1975 Conference on Prevention and Control of Oil Pollution". American Petroleum Institute, Washington, D.C., 1975, p 115. (9) 0. Deshimaru, Bull. Jpn. SOC. Sci. Fisheries, 37, 297 (1971). (10) W. E. May, S. N. Chesler, S. P. Cram, B. H. Gump, H. S. Hertz, D. P. Enagonio, and S. M. Dyszel, J . Chromatogr. Sci., 13, 535 (1975). (11) S. N. Chesler, E. H. Gump, H. S. Hertz, W. E. May, S. M. Dyszel, and D. P. Enagonio, Abtl. Bur. Stand. (U.S.),Tech. Note. No. 889, Washington, D.C., 1976, 73 pp. (12) H. Hachenberg and A. P. Schmidt, "Gas Chromatographic Headspace Analysis", Heyden and Sons Ltd., London, 1977, pp 10-18. (13) S. A. Wise, S. N. Chesler, H. S. Hertz, L. R. Hiipert, and W. E. May, Anal. Chem., 49, 2306 (1977).
RECEIVEDfor review November 29, 1977. Accepted February 13, 1978. The authors acknowledge partial financial support from the Bureau of Land Management through interagency agreement with the National Oceanic and Atmospheric Administration, under which a multi-year program responding to needs of petroleum development in the Alaskan continental shelf is managed by the Outer Continental Shelf Environmental Assessment Program (OCSEAP) Office. Paper presented a t the 172nd American Chemical Society National Meeting, San Francisco, Calif., August 1976. Identification of any commercial product does not imply recommendation or endorsement by the National Bureau of Standards, nor does it imply t h a t the material or equipment identified is necessarily the best available for the purpose.
