Envlron. Scl. Technol. 1983, 17, 548-553
expect a complete oxidation of the P A H s within this reaction time, provided no faster reacting compounds are present. The rate constants also may be used for the prediction of half-lives of PAHs in the lower atmosphere. It is now generally accepted that, at least at low altitudes and in regions of moderate and high humidity, the atmospheric aerosols consist of liquid shells surrounding the opaque nucleus (13). The aerosol water content at humidities above 40% is estimated to be at least 30% of the total particle weight (14). This property permits the chemistry of atmospheric aerosols to be treated as in aqueous solution (13). Due to their low vapor pressures the PAHs released into the atmosphere are associated with aerosols and airborne particulate matter. With a steady-state concentration of tropospheric ozone of 2 X lo4 M in clean air (15), the half-life values in air are 2.5 h for pyrene, 6 h for phenanthrene, and 14 h for benzo[a]pyrene. This view is consistent with the observation recently made by Pitts et al. (16) and Katz et al. (17)that benzo[a]pyrene and other PAH’s react in the adsorbed state within hours in the dark in ozone-containing atmosphere to yield oxidized derivatives of PAH’s. ’RegistryNo. Pyrene, 129-00-0; phenanthrene, 85-01-8; benzo[a]pyrene, 50-32-8.
Literature Cited (1) Gelboin, H. V., Ts’O, P. 0. P., Eds. “Polycyclic Hydrocarbons and Cancer”; Academic Press: New York, 1978; VOl. 1, 2. (2) Neff, J. M. “Polycyclic Aromatic Hydrocarbons in the Aquatic Environment”; Applied Science: London, 1979. (3) Reichert, J. GFW, Gas-Wasserfach 1969, 110, 477-480. (4) Gomella, C. J . Am. Water Works Assoc. 1972,64,39-45.
(5) Il’nitakii, A. P.; Khesina, A. Y.; Cherkinskii, S. N.; Shabad, L. M. Gig. Sanit. 1968, 33, 8-13; Hyg. Sanit. 1968, 33, 323-327. (6) Radding, S. B.; Mill, T.; Gould, C. W.; Liu, D. H.; Johnson, H. L.; Bomberger, D. C.; Fojo, C. V. US Environmental Protection Agency, PB-250948, Washington, DC, 1976. (7) Andelman, J. B.; Snodgrass, J. E. CRC Crit. Rev. Enuiron. Control 1974, 4 , 69-83. (8) Harrison, R.M.; Perry, R.; Wellings, R. A. Water Res. 1975, 9, 331-346. (9) Braunstein, H. M.; Copenhaver, E. D.; Pfuderer, H. A. “Environmental, Health and Control Aspects of Coal Conversion: An Informative Overview”; ORNL/EIS-94, 1977; VO~.1, pp 4-136. (10) Hoigng, J.; Bader, H.Water Res. 1983, 17, 173-183. (11) Sforzolini, G.S.; Savino, A.; Monarca, 6. Ig. Mod. 1974,66, 595-619. (12) Burleson, G. R.; Caulfield, M. J.; Pollard, M. Cancer Res. 1979, 39, 2149-2154. (13) Graedel, T. E.; Weschler, C. J. Rev. Geophys. Space Phys. 1981,19,505-539. (14) Friedlander, S. K. Environ. Sci. Technol. 1973, 7, 235-240. (15) Levy, H. Science (Washington, D.C.) 1971,173,141-143. (16) Pitts, J. N., Jr.; Lokensgard, D. M.; Ripley, P. S.; Van Cauwenberghe, K. A.; Van Vaeck, L.; Shaffer, S. D.; Thill, A. J.; Belser, W. L., Jr. Science (Washington,D.C.) 1980, 210, 1347-1349. (17) Katz, M.; Chan, C.; Tosine, H.; Sakuma, T. “Polynuclear Aromatic Hydrocarbons”; Jones, P. W., Leber, P., Eds.; Ann Arbor Science: Ann Arbor, MI, 1979; pp 171-189.
Received for review July 26,1982. Revised manuscript received February 7,1983. Accepted April 7,1983. This work was performed on the basis of a German-Yugoslav scientific cooperation program. The financial support by the International Biiro, Kernforschungsanlage Jiilich, and by the Republic Council for Science of Croatia (SZZ In is gratefully acknowledged.
Preparation of Aqueous Petroleum Solutions for Toxicity Testing Kjetill Dstgaard and Arne Jensen” Institute of Marine Biochemistry, University of Trondheim, N-7034 Trondheim NTH, Norway
Direct fluorescence spectroscopy revealed that 5-10 days of slow stirring in a closed system was needed to reach apparently saturated water-soluble fractions (WSFs) of Ekofisk crude oil in seawater, provided the process was carried out in complete darkness. Under normal laboratory illumination no saturation of the naphthalene fraction was reached within 3 weeks. Stronger light increased the concentration of the naphthalene fluorescing material. The fluorescence levels reached after 10 days of standardized stirring under strictly defined light conditions were reproducible within 6%. Chemical analysis (GS/MS) of the extracts was in accordance with the fluorescence data. The toxicity of the WSF to the diatom Skeletonema c o s t a t u m was closely related to the recorded fluorescence, following the changes induced by illumination. Growth inhibition was found to depend on initial algal biomass. It is concluded that the stirring times and the variable light conditions commonly used cannot give reproducible WSFs for toxicity testing. Furthermore, standardization of initial algal biomass seems to be required in quantitative studies of petroleum toxicity to microscopic algae. Introduction
The toxic effects of crude oil and petroleum products in the marine environment have been extensively studied 548
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(cf. ref 1-3) and continue to attract considerable interest (4-6). The studies are usually based on what is regarded as true solutions of petroleum compounds in seawater, the so-called water-soluble fractions (WSFs). These are normally prepared either by slow stirring of oil on water (7-9) or by vigorous shaking of the two components, followed by phase separation when no oil droplets are visible in the water (10-12). Shaw and Reidy (13) have shown that slow stirring with no formation of visible oil droplets gave an aqueous phase that was highly enriched in phenols and aromatic hydrocarbons, relative to the composition of the oil. Contrary to this, vigorous shaking gave extensive emulsification, and the lipid component in the water was very similar to the original oil. Besides the mixing mode other experimental factors such as duration of treatment, temperature, light, and microbial activity will influence the physical nature and chemical composition of the extract. No general standardization of the procedures has been introduced, and the oil extracts that have been produced must have varied accordingly in nature and composition. The complexity of oil composition and behavior prohibits a complete chemical description of oil dispersions (14). Advanced techniques involving chromatographic separation and mass spectrometry are too expensive to be
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0 1983 American Chemical Society
used extensively in routine testing. In this situation an acceptable compromise seems to involve rigid standardization of the preparation of aqueous oil solutions for toxicity testing and the use of a simple screening method to follow the process and to guide the sampling for more thorough chemical analysis. Batch culture systems are commonly used in toxicity studies because of their experimental simplicity. From a biological point of view they represent a very complex situation, where the changes in actual dose and growth conditions are unknown. Additional problems may therefore arise when such systems are used to assess the toxicity of aqueous petroleum solutions. The present paper describes our attempts to prepare reproducible solutions of crude oil in seawater and the use of direct fluorescence spectroscopy to monitor the process. It also reports our attempts to relate the fluorescence of the extracts to their phytotoxicity observed in batch cultures. Experimental Section The seawater used in this study was taken from a depth of 30 m in a practically unpolluted area, filtered, diluted to 25%0salinity, enriched in nutritional components to the f/10 level of the culture medium of Guillard and Ryther (I5),and sterilized in an autoclave. Standard temperature during preparation and testing was 14 "C. Illumination was supplied in a regular day-night cycle of 14:lO h and measured by a Li-Cor cosine-corrected quantum sensor (unidirectional) or a Biospherical Instruments Inc. QSL100 meter (total spherical distribution), both with spectral range 400-700 nm. Solutions of Ekofisk crude oil were prepared by carefully layering one part of oil (200 mL) on top of 20 parts of sterile seawater (4 L) held in a 5-L glass bottle and closed with a silicone stopper. By use of a standard magnetic stirrer run at reduced voltage, gentle stirring was achieved, i.e., no deformation of the oil-water interface was observable. Samples were taken by pumping through a glass/silicone rubber tubing located below the level of the oil layer and collected in sterile brown glass medical flasks that were completely filled before closing. Direct fluorescence spectroscopy of the samples was applied for monitoring the dissolution process. Emission spectra a t excitation wavelength 230 nm, favoring the fluorescence of naphthalenes around 335 nm, and at excitation wavelength 265 nm, favoring the fluorescence of phenols around 300 nm (161, were recorded in a PerkinElmer 3000 fluorescencespectrometer. Figure 1illustrates typical spectra and the recorded values designated "naphthalene fraction" and "phenol fraction" here. The fluorescence unit was arbitrarily chosen and is the same in all experiments. The chemical content of highly volatile components was determined by head-space analysis (17). Briefly, the compounds were stripped from the solution at 60 "C by a stream of nitrogen and adsorbed on active charcoal in a capillary. This was then placed in the injector of a gas chromatograph and analyzed. The components were identified by coupled gas chromatography/mass spectrometry (GC/MS) and quantified by using benzene-d6as internal standard. Less volatile compounds were extracted with dichloromethane (17). The content of components with boiling point above that of toluene (C7+fraction) was determined by gas chromatography (packed column), calibrated against known amounts of Ekofisk crude. Single components of the same extract were identified by GC/MS and quantified by use of the internal standards diphenyl-d,, and anthracene-d,, ( I 7).
A
Naphthalene fraction a, C
U
E
-a U
Phenol fraction
Wave length inrn)
Figure 1. Naphthalene fraction and phenol fraction defined by direct fluorescence. Characteristic emission spectra for a solution of Ekofisk crude oil in seawater (E) and for the seawater used (sea): (A) exitation wavelength 230 nm; (B) Exitation wavelength 265 nm.
Skeletonemu costatum (Grev.) Cleve, clone Skel-5 (unialgal) isolated by S. Myklestad, Institute of Marine Biochemistry, Trondheim, Norway, was kept as a stock culture at 14 OC and approximately 70 peinsteins/(m2 s) of light in a day-night cycle of 14:lO h. Experiments were performed under the same conditions in closed 250-mL Erlenmeyer flasks containing 100 mL of seawater. Exponentially growing stock cultures were concentrated by centrifugation before inoculation of 1mL per experimental culture, giving a cell density of approximately 8 X lo4 cells/mL unless otherwise stated. Growth was followed for 4 days by daily sampling with a syringe through a silicone rubber cap over a short side arm located below the surface of the medium. The cells were counted in a hemocytometer and in vivo fluorescence was measured in a Turner 111fluorometer. The increase in fluorescence after addition of 3-(3,4-dichlorophenyl)-l,l-dimethylurea (DCMU) was also recorded to monitor the photosynthetic capacity (18). Results Studies of the Dissolution Process. Preliminary experiments had shown that no stable fluorescence level could be achieved within 3 weeks (16) when the WSF of Ekofisk crude oil was prepared by slow stirring under normal indoor illumination. Figure 2A illustrates the kinetics of the dissolution process performed in complete darkness. Although the content of phenols in the water phase apparently reached equilibrium after 2 days, the naphthalene fluorescence showed a small but significant increase throughout the latter part of the experiment. Compared with the results obtained under exposure to light (16),however, the naphthalene fraction can be considered relatively constant. Parts B and C of Figure 2 illustrate the influence of light intensity on the preparation of WSF. A defined phenol fluorescence was reached in all cases, but the final level was dependent on the light intensity. Contrary to this, the naphthalene fraction showed persistent increase with time for the total period of 10 days. This increase was strongly enhanced at the higher light intensity. The effect seemed to be cumulative. Although the general shape of the curves in Figure 2 was reproducible, the development during the first 2-3 days was not. This was probably related to small variations in the stirring rate. The fluorescence values obtained after Environ. Sci. Technol., Vol. 17, No. 9, 1983 549
A
h
1000
looOL Time (days1
I
I)
500
Flgure 3. Behavior of aqueous solutions of Ekofisk crude oil under laboratory conditions. Fluorescence of naphthalene fraction (n) and phenol fraction (p) as a function of time: (A) preparation in complete darkness disturbed by a 14-h pulse (limited by broken lines) of strong light (730 peinsteins/(m2s)); (E) storage in the absence of the oil layer; closed system; illumination 570 peinsteins/(m2s).
,~._---.*-.
,'
:;
' I
0 4
5
6
9
10
17
T i m e (days1
Flgure 2. Preparation of aqueous solutions of Ekofisk crude oil by slow stirrlng under laboratory conditions. Fluorescence of naphthalene fraction (n) and phenol fraction (p) recorded as a function of time in (A) total darkness, (E) 130 peinsteins/(m* s)(total spherical distribution 14 h light: 10 h dark), and (C) 570 peinsteins/(m2s) (total spherical distribution 14 h light: 10 h dark).
Table I. Fluorescence, in Relative Units (See Text), of Aqueous Solutions of Ekofisk Crude Oil Stirred for 10 Days under Different Light Levels in 14 h Light:lO h Dark Regimesa light, ueinsteins / (m* s) in distilled water vert in seawater naphth phenol total comspher- ponaphth phenol frac- fracical nent fraction fraction tion tion 0 0 1257(23) 824(21) 1765 1070 70 20 1361(40) 8 0 5 ( 1 2 ) 70 1528(89) SOS(32) 130 570 360 1 8 1 3 ( 4 8 ) 697 (40) 2310 920 a Figures in brackets are standard deviations based on 3-4 indeDendent exDeriments.
2-3 days and the related oil contents were significantly lower than those reached after prolonged stirring. We therefore expect the stirring times of 12-24 h generally used (see, e.g., ref 7,8,19)in the preparation of WSF to be too short to secure reproducible extracts. The reproducibility in fluorescence observed after longer stirring times is indicated in Table I, giving data for 10 days of stirring. The standard deviations were within *6% for each of the four different light regimes tried. This is close to the accuracy of the fluorescence method itself (16), loss of volatiles during sample handling being a major source of error. The photochemical reactions leading to the results of Figure 2 and Table I are not known. As shown in Table I, oil on distilled water behaved in a similiar way, the only difference being the expected increase in solubility of the oil (20). 550
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Table 11. Chemical Composition of Seawater Solutions of Ekofisk Crude Oil Stirred for 1 0 Days under Different Light Levels in 14 h Light:lO h Dark Regimes
compound/fraction Components lighter than benzene benzene toluene C, fraction ethylbenzene xylenes C, alkylbenzenes naphthalene methylnaphthalenes methylbenzaldehyde phenol C ,-C, a1k ylphenols total a Not detected.
complete darkness 4 PPm 6P P ~
1 PPm 5.7 ppm 23 PPb 270 ppb 72 PPb 43 ppb 37 ppb nda 3 ppb 214 ppb 17 ppm
570 ueinsteins / ( m Zs) 5.5 ppm 8P P ~
1.3 ppm 9.1 ppm 1 7 ppb 128 ppb 47 ppb 130 ppb 41 PPb 3 PPb nd 130 ppb 24 PPm
The connection between illumination and increase of the naphthalene fraction seemed to be indirect, as illustrated in Figure 3A. Here, slow stirring was applied in complete darkness, until a stable fluorescence level was reached (9-10 days). This was followed by a 14-pulse of strong light. The increase in naphthalenes started only toward the end of this period and became prominent in the following 2 days, giving a new stable level (Figure 3A). This treatment led to a slight reduction in the fluorescence of phenols (Figure 3A). Illumination of a crude oil solution in the absence of an overlaying oil layer rapidly reduced the fluorescence of the naphthalene fraction (see Figure 3B). Similar kinetics were observed when pure naphthalene solutions were illuminated (results not included). Figure 3B indicates how variable the dose may be when the toxicity of oil is tested in closed-batch systems exposed to light. Chemical Composition. The results of the GC/MS analysis have been summarized in Table 11. A more complete list of components found in aqueous solutions of Ekofisk crude oil has been given elsewhere (22). As expected for closed systems, highly volatile compounds like benzene and toluene composed a major fraction (Table 11).
A
5
K l T
0.5
1
Retention time
----j
Figure 4. Gas chromatograms of aqueous solutions of Ekofisk crude oil stirred for 10 days under different conditions. Naphthalene (n) and methylnaphthalenes (mn) are indicated together with the internal and anthracene-d, (IS2):(A) complete standards biphenyl-d,, (IS,) darkness; (B) illumlnated at 570 (heinsteins/(mB s).
Since these compounds are easily lost during handling of samples, the observed differences between the two preparations are not regarded as significant. The photoinduced increase in the content of compounds with boiling point higher than toluene (C7+fraction) was, on the other hand, quite clear and consistent. A more moderate illumination (130 peinsteins/(m2 s)) gave a C7+fraction (8.7 ppm) between that produced in the dark (5.7 ppm) and that found in strong light (9.1 ppm). Only a few of the compounds within the C7+fraction have been identified and quantified. The identified naphthalenes increased their concentrations upon illumination, while phenol and its alkyl derivates decreased. These results are qualitatively in accordance with the fluorescence data of Table I. The corresponding gas chromatograms given in Figure 4 illustrate a dominating increase in heavier compounds upon illumination. These compounds could not be identified by GC/MS due to their partial instability during the run (17). The pronounced enrichment in aromatic compounds in the water phase is in accordance with results obtained by others for crude oil solutions prepared by slow stirring (13, 22). Toxicity. Growth of Skeletonema costaturn in closedbatch cultures containing identical loads of aqueous crude oil solutions was followed for 4 days after inoculation of different cell concentrations. As shown in Figure 5A, the toxic effects were highly dependent on the initial cell density. Although a similar dependency has been observed by others for the toxicity of heavy-metal ions (23),it has not been taken into account when evaluating the phytotoxicity of crude oils. Additional problems arise because the biomass per cell of diatoms may vary considerably due to partial synchrone in auxospore formation (24). Ex-
Time (days) Flgure 5. Growth of Skelefonerna costaturn in the presence of aqueous solutions of Ekofisk crude oil: cultures with completely blocked photosynthesis (D);viable cultures (e). Control cultures are marked R. (A) Influence of initial cell density (lo4ceilslml) as indicated; (B) influence of dilution of the standard solution (percent of undiluted sample) as indicated on curves; (C) influence of the length of the stirring period (days) as indlcated; (D)influence of illumination during stirring (peinsteins/(m2 s) in a 14 h light: 10 h dark regime) as indicated.
periments (not included) have indicated that it is the initial biomass that should be standardized in order to obtain reproducible results. In the most critical sense the independent experiments in Figure 5 are therefore not quantitatively comparable. Figure 5B illustrates that a crude oil solution (obtained by stirring for 10 days under moderate illumination) killed the algal culture after 2 days of exposure. Dilution to 25% of original strength restored the growth to the control level after a lag period of 1 day. Environ. Sci. Technol., Vol. 17, No. 9, 1983 551
As seen from Figure 5C the toxicity increased steadily with the period of stirring under illumination. It is of particular interest that the extract obtained after 5 days gave zero growth, while the 10-day sample completely killed the culture within 2 days (Figure 5C). This should be compared to the corresponding increase in fluorescence seen in Figure 2B. No saturation in toxicity was observed for the crude oil solutions that were exposed to light during stirring. Figure 5D shows how the toxicity of the aqueous extracts increased with increasing light intensity. Together parts C and D of Figure 5 indicate that the toxic compounds accumulated during the treatment and that total integrated illumination appeared to determine the photoinduced phytotoxicity of crude oils (25,26). Although Figure 5,C and D, might be taken to suggest a correlation between fluorescence of naphthalenes (Figure 2; Table I) and toxicity, it must be remembered that most of the photoinduced changes in composition could not be identified (see Figure 4). Discussion
The preparation of defined doses seems to be one of the major obstacles in quantitative studies of the toxicity of petroleum to aquatic organisms. The commonly used methods involve true dissolution, emulsification, and transformation of compounds by photooxidation. True dissolution is a selective process leading to enrichment in the water phase of the more polar compounds of the original oil (13). The final dose of such compounds occurring in very low quantities in the oil will vary with the proportion of oil to water. Emulsification does not involve selection and gives droplets of the original oil dispersed in the water phase in addition to compounds in true solution (13). It is unfortunate that the designation “water-solublefraction” is also being used in the latter case. Earlier studies (13,16) have shown that stirring has to be very gentle to avoid formation of dispersion. Photooxidation of crude oil leads to polar compounds that are selectively extracted into the water. This may influence both the solubility properties and the toxicity of the aqueous phase (25,26). In the present study direct fluorescence spectroscopy (16) was used to follow the dissolution process. This screening method detects only part of the total oil components in the water, mainly aromatic compounds. Solutions with identical fluorescence spectra may therefore still be quite different in composition. On the other hand difference in spectral properties proves real difference in chemical composition of such solutions. The high sensitivity of the method to small variations in the conditions during preparation of solutions shows its practical usefulness. Furthermore, the conclusions reached on the basis of the spectroscopic data were supported by chemical analysis (Table 11). It is quite clear, however, that many polar compounds remained undetected in both the fluorometric and the chemical analyses. Only when comparisons are restricted to samples prepared under one set of rigorously standardized conditions do we expect fluorescence data to reflect the general reproducibility of the preparation procedure (cf. Table I). Fluorescence spectroscopy must therefore be followed by adjoined toxicity studies before conclusions can be drawn. According to the fluorescence studies, stirring had to continue for 5-10 days in complete darkness to give apparently equilibrated solutions (see Figure 2). This is much longer than the 12-24 h normally used (2,6-8,11,19,27, 28,and others). The concentrations reached after 1-2 days were significantly lower and very dependent upon small 552
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variations in stirring rate and equipment configuration, In the light similar kinetics were observed for the first several days, but no saturation of the water phase with oil components was attained within 3 weeks (Figure 2). This increase in fluorescence with time was closely paralleled by enhanced toxicity of the extracts to the test alga (Figure 5C). According to Table 11reproducible extracts could still be produced after 10 days of slow stirring under strictly standardized conditions (including light conditions). When light intensity was changed instead of the time of exposure, the increase in fluorescence (Figure 2; Table 11) was also followed by increased toxicity (Figure 5D). Whether this effect was caused by the elevated content of oil components in general or by special phytotoxicity of the oxidation products cannot be decided. It has been reported (25) that photooxidation will increase the phytotoxicity of crude oil, and Figure 5D shows that this effect was significant even under normal laboratory illumination. This has often been overlooked, although Giddings and Washington (19) have pointed out the importance of screening against light when preparing aqueous petroleum solutions. Due to inherent differences in photochemical response, any toxicity ranking of oils and petroleum products is expected to depend on the light conditions during the test. Preparation and testing of aqueous oil extracts in complete darkness seems rather unnatural and cannot be resorted to for photosynthetic organisms. Performance of a series of tests under a set of well-defined light conditions appears to be a better alternative. Another problem that has been frequently overlooked is the influence of biomass reflected in Figure 5A. For macroscopic organisms toxicity data always refer to the weight of the organism (i-e.,units of toxin per unit of body weight). In the case of microorganisms the concentration of the toxic substance that produces the response is determined without direct reference to the biomass of the organimsm. It is known that biomass is important in the determination of the toxicity of heavy metals to microalgae (23). The effect is explained as a result of uptake or binding of the toxic compounds. This lowers the concentration (activity) of the toxic material in the medium, thereby enabling surviving cells to grow more or less normally. A similar mechanism may be operating also for lipophilic toxins in water. The relationship between solubility and toxicity established by Hutchinson et al. (29) as well as the steep slope of the dose-response curve indicated in our data (Figure 5B) may be understood in light of this hypothesis. The neglection of methodological problems involved reduces the value of the rapidly growing literature on the biological effects of petroleum components on aquatic organisms. Appreciation of the technical problems and rigid standardization of the procedures appear to be essential. It is hoped that the approach presented here could also prove generally useful in studies of other common experimental systems. Acknowledgments
We thank A. Andrewes for skillful experimental assistance and F. Oreld and R. G. Lichtenthaler at the Central Institute of Industrial Research, Oslo, for the chemical analysis performed. Registry No. Benzene, 71-43-2; toluene, 108-88-3; ethylbenzene, 100-41-4; xylene, 1330-20-7;naphthalene,91-20-3;methylbenzaldehyde, 1334-78-7;phenol, 108-95-2.
Literature Cited (1) Corner, E. D. S. Adv. Mar. Bid. 1978, 15, 289.
Envlron. Sei. Technol. 1983, 17, 553-555
(2) Anderson, J. W.; Neff, J. M.; Cox, B. A.; Tatem, H. E.; Hightower, G. M. Mar. Biol. 1974, 27, 75. (3) Mironov, 0. G.; Shchekaturina, T. L.; Tsimbal, I. M. Mar. Ecol. Prog. Ser. 1981,5, 303. (4) Maher, W. A. Bull. Environm. Contam. Toxicol. 1982,29, 268. (5) Bastian, hl.V.; Toetz, D. W. Bull. Enuironm. Contam. Toxicol. 1982, 29, 531. (6) Widdows, J.; Bakke, T.; Bayne, B. L.; Donkin, P.; Livingstone, D. R.; Lowe, D. M.; More, M. N.; Evans,S. V.; More, S. L. Mar. Biol. 1982, 67, 15. (7) Kauss, P.; Hutchinson, T. C.; Soto, C.; Hellebust, J.; Griffiths. M. Proc. J. Conf. Prev. Control Oil S d l s 1973, 703. (8) Pulich, W. M., Jr.; Winters, K.; Van Baalen, C. Mar. Biol. 1974, 28, 87. (9) Mahoney, B. M.; Haskin, H. H. Enuironm. Pollut., Ser. A 1980, 22, 123. (10) Prouse, N. J.; Gordon, D. C., Jr.; Keizer, P. D. J. Fish. Res. Board Can. 1976, 33, 810. (11) Soto, C.; Hellebust, J.; Hutchinson, T. C.; Sawa, T. Can. J . Bot. 1976, 53, 109. (12) Kusk, K. 0. Bot. Mar. 1980,23, 587. (13) Shaw, D. G.; Reidy, S. K. Environ. Sei. Technol. 1979,13, 1259. (14) Petrakis, L., Weiss, F. T., Eds. “Petroleum in the Marine Environment”; papers from a symposium, Miami Beach, FL, Sept 1978; American Chemical Society: Washington DC, 1980. (15) Guillard, R. R. L.; Ryther, J. H. Can. J. Microbiol. 1962, 8, 229. (16) Ostgaard, K.; Jensen, A. Int. J. Environ. Anal. Chem. 1982, 14, 55.
(17) Berg, N.; Gustavsen, K.; Gjcls, N.; Lichtenthaler, R. G.; Oreld, F.; Vadum, K.; Ofsti, T. “Chemical Analysis of Water Soluble Petroleum Fractions and Studies of Their Aceumulation in Flounders”; SI Report 780702-1: Oslo, 1980. (18) Samuelsson, G.; Oquist, G. Physiol. Plant. 1977, 40, 315. (19) Giddings, J. M.; Washington, J. N. Enuiron. Sei. Technol. 1981, 15, 106. (20) Price, L. C. Am. Assoc. Pet. Geol. Bull. 1976, 60, 213. (21) Ostgaard, K.; Eide, I.; Jensen, A. Mar. Environ. Res., in press. (22) Winters, K.; ODonell, R.; Batterton, J. C.; Van Baalen, C. Mar. Biol. 1976, 36, 269. (23) Steeman Nielsen, E.; Wium-Andersen, S. Mar. Biol. 1970, 6, 93. (24) Migita, S. Bull. Jpn. SOC.Sei. Fish. 1967, 33, 392. (25) Lacaze, J. C.; Villedon de Naide, 0. Mar. Pollut. Bull. 1976, 7,73. (26) Larson, R. A.; Bott, T. L.; Hunt, L. L.; Rogenmuser, K. Environ. Sci. Technol. 1979, 13, 965. (27) Kauss, P. B.; Hutchinson, T. C. Environ. Pollut. 1975,9, 158. (28) Soto, C.; Hellebust, J. A.; Hutchinson, T. C. Can. J. Bot. 1979,24, 2717. (29) Hutchinson, T. C.; Hellebust, J. A,; Mackay, D.; Tam, D.; Kauss, P. In “Proceedings-1979 Oil Spill Conference”; American Petroleum Institute: Washington DC, 1979; p 541.
Received for review July 29,1982. Revised manuscript received March 22,1983. Accepted April 6, 1983. This work is part of the Norwegian Marine Pollution Research and Monitoring Program.
On the Formation of Mutagens in the Chlorination of Humic Acid Knut P. Kringstad, Pierre 0. Ljungqulst, Filipe de Sousa, and Lars M. Stromberg” Swedish Forest Products Research Laboratory, Box 5604, S-114 86 Stockholm, Sweden
The present investigation shows that the strong direct-acting mutagens 1,3-dichloroacetone and 2-chloropropenal are formed at low levels in the chlorination of humic acid. These results therefore suggest that these two compounds may also possibly contribute to the mutagenic activity of chlorinated drinking water. Introduction A large number of organic compounds have been identified as constituents of chlorinated drinking and wastewater (1-4). Several of these compounds were found to be mutagens and/or carcinogens (5-7). The compounds may be present in the raw water itself but also may be produced in the water chlorination process by reactions between chlorine and organic matter such as humic and fulvic acids (1, 8-11). The presence of mutagens and carcinogens in drinking water has caused concern and led to considerable efforts to identify and characterize as many as possible of the organic compounds present. Recently, we described the identification and mutagenic properties of some chlorinated aliphatic compounds present in the spent liquor from the chlorination of softwood kraft pulp (12,13). Several mutagenic compounds were found, and some belong to those previously identified in chlorinated drinking water. This is not surprising, since the chemical structures of humic and fulvic acids on one hand and of residual lignin in kraft pulp on the other are related (14). However, two of the mutagens in the spent chlorination liquor have so far not been identified in 0013-936X/83/09 17-0553$0 1.5010
drinking water. These direct-acting mutagens, which appeared to be particularly strong, were 1,3-dichloroacetone and 2-chloropropenal. The latter very likely carries a major responsibility for the total mutagenic activity (Ames test, Salmonella typhimurium TA 1535) of the spent chlorination liquor (13). This paper describes studies carried out with the primary objective of determining to what degree, if any, 1,3-dichloroacetoneand/or 2-chloropropenal are formed in the chlorination of humic acid and of determining its consequent potential contribution to drinking water mutagenicity. Experimental Section
Humic Acid Materials. Four humic acid materials were studied. Two of these were isolated from aquatic sources, and two were commercially available samples. Isolation procedures and characteristics for the various materials were as follows: Aquatic Sample I: Surface water (1300 L) was collected from a lake in the southern part of Sweden. The permanganate number of the water was 67 mg of KMn04/L. Humic acid was isolated from the water by acidification with HCl to pH 2.1, following a lengthy previously described procedure (15). Elemental analysis of the material revealed a C:H:O:N:S ratio of 1.00:1.20:0.48:0.05:0.005. The ash content was found to be 13%. The IR absorption spectrum showed absorption bands at -3400 (br), 2920, -1710,1620 (slightly broadened), 1525 (w), 1450-1350 (w), and 1080-1030 cm-’ (weak)
0 1983 American Chemical Society
Environ. Sci. Technol., Vol. 17, No. 9, 1983 558
