(27) (28) (29) (30)
(38) Wakeham, S. G. Woods Hole Oceanographic Institution, Woods Hole, Massachusetts, unpublished data, 1983. (39) Jordan, R. E.; Payne, J. R. In “Fate and Weathering of
imental Ecosystems”; Grice, G. D. Reeve, M. R., Eds.; Springer Verlag: New York, 1982; pp 81-95. Santschi,P. H.; Carson, S.; Li, Y.-H. In “Marine Mesocosms: Biological and Chemical Research in Experimental Ecosystems”; Grice, G. D., Reeve, M. R., Eds.; Springer Verlag: New York, 1982; pp 98-109. Wakeham, S. G.; Goodwin, J. T.; Davis, A. C. Woods Hole Oceanographic Institution Technical Report 82-36,1982; p 62. Bopp, R. F.; Santschi, P. H.; Li, Y.-H.; Deck, B. L. Org. Geochem. 1981,3,9-14. Broecker, W. S.; Peng, T.-H. Tellus 1974, 26, 21-35. Liss, P. S.; Slater, P. G. Nature (London) 1974, 247,
Petroleum Spills in the Marine Environment”; Ann Arbor Science Publishers: Ann Arbor, MI, 1980; p 174. (40) Claus, D.; Walker, N. J. Gen. Microbiol. 1964,36,107-122. (41) Gibson, D. T.; Koch, J. R.; Kallio, R. E. Biochemistry 1968, 7, 2653-2662. (42) Gibson, D. T.; Gschwendt, B.; Yeh, W. K.; Kobal, V. M. Biochemistry 1973, 12, 1520-1528. (43) Gibson, D. T.; Koch, J. R.; Kallio, R. E. Biochemistry 1968, 7, 3795-3802. (44) Alexander, M. Adv. Appl. Microbiol. 1965, 7, 35-80. (45) Grob, K.; Grob, G. J. Chromatogr. 1974, 90, 303-313. (46) Lee, R. F. In “Proceedings, 1977 Oil Spill Conference
181-184. (31) Schwarzenbach, R. P.; Molnar-Kubica, E.; Giger, W.; Wakeham, G. Enuiron. Sei. Technol. 1979,13,1367-1373. (32) Nixon, S. W.; Alonso, D.; Pilson, M. E. Q.; Buckley, B. A.
s.
In “Microcosms in Ecological Research”; Glessey, Ed.; DOE Symposium Series CONF-F81101, National Technical Information Service: Washington, D.C., 1980; pp 818-849. (33) Bouwer, E. J.; Rittmann, B. E.; McCarty, P. L. Environ. Sci. Technol. 1981, 15, 596-599. (34) Ballschmitter, K.; Scholz, C. Chemosphere 1980,9,457-467. (35) Marinucci, A. C.; Bartha, R. Appl. Enuiron. Microbiol. 1979, 38, 811-817. (36) Schwarzenbach, R. P.; Westall, J. C. Enuiron. Sei. Technol. 1981,15, 1360-1367. (37) Schwarzenbach, R. P. Swiss Federal Institute for Water
Resources and Water Pollution Control (EAWAG), private communication, 1982.
(Prevention, Behavior, Control, Cleanup)”; American Petroleum Institute Public8tion 4 2 M Washington, D.C., 1977; pp 611-616. (47) Leo, A.; Hansch, C.; Elkins, D. Chem. Rev. 1971, 71, 525-616.
Received for review January 7,1983. Revised manuscript received May 19,1983. Accepted May 24,1983. This investigation was supported by U S . Environmental Protection Agency Grants R80607212 and CR807795 to the Marine Ecosystems Research Laboratory (MERL),University of Rhode Island, and subcontracted to WHOI, and National Oceanic and Atmospheric Administration, Office of Marine Pollution Assessment, Grant NA81RAD00015.
Biodegradation of the Monochlorobiphenyls and Biphenyl in River Water Robert E. Bailey,” Stanley J. Gonsior, and Wayne L. Rhinehart
Environmental Sciences Research Laboratory, Dow Chemical U.S.A., Midland, Michigan 48640 The rates of primary biodegradation, as well as mineralization, of the three isomers of monochlorobiphenyl (MCB), biphenyl (BP), and 2,2’,4,4’-tetrachlorobiphenyl (TCB) have been determined by using a river water dieaway test. Analysis of river water extracts by high-performance liquid chromatography showed the times for 50% degradation of parent MCBs were 2-3 days and 1.5 days for BP at initial concentrations of about 1 pg/L. The unchlorinated ring degraded initially and after a 50-day incubation, approximately 50% of the theoretical I4CO2 was recovered from the subsequent degradation of the chlorinated ring of the MCBs. The rate af biodegradation of higher concentrations was somewhat slower. No degradation of TCB was observed over 98 days under conditions of this test. H
Introduction The biodegradation of the monochlorobiphenyls (MCBs) has been reported by several groups in studies conducted both with pure cultures (1,2) and by mixed populations (3-8).A comparison of the rate of MCB biodegradation with the rates of biphenyl (BP) and 2,2’,4,4’-tetrachlorobiphenyl (TCB) allows an estimation of the environmental importance of their biodegradation. TCB is a typical component of the polychlorinated biphenyls (PCBs) found in environmental samples. TCB has been observed to not degrade under conditions where other PCB congeners were degraded (1,9). In contrast, BP is widely distributed in the environment as a component of petroleum and as an article of commerce but was observed only at concentra0013-936X/83/0917-0617$01.50/0
tions of 0.1-0.5 pg/L in natural waters (10). The production of chlorobenzoic acid (CBA) from MCB biodegradation has been reported several times (1,2,11). Recently, (2-chlorobenzoyl)formicacid was reported as a metabolite of 2-MCB (5). The CBAs are well-known to be biodegradable, although often slowly or with a lag period depending on the bacterial inoculum used (12-21). The purpose of this study was to determine the rate of primary biodegradation as well as mineralization of all the MCB isomers for use in predicting their environmental removal rate. Each of the three MCB isomers was studied separately to observe any isomer effects and compared with BP and TCB.
Experimental Procedures Chemicals. Radioactively labeled MCBs (uniformly 14C labeled on the chlorinated ring) were synthesized by Pathfinder Laboratories (St. Louis, MO). The radiochemical purity was determined by reverse-phase highperformance liquid chromatography (HPLC) and liquid scintillation counting. The radiochemical purity and specific activity of the products were the following: 2MCB, 94%, 11.84 mCi/mmol; 3-MCB, 91%, 14.28 mCi/ mmol; 4-MCB, 87%, 18.05 mCi/mmol. BP (The Dow Chemical Co.) and TCB (Pathfinder Laboratories) were uniformly 14C labeled with radiochemical purities and specific activities of 99%, 6.647 mCi/mmol and 99%, 11.46 mCi/mmol, respectively. River Die-Away Studies. River water was obtained from the Tittabawassee River, Midland, MI. The sampling site was upstream from any significant industrial activity
0 1983 American Chemical Society
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or municipal discharge. The Tittabawassee River drains primarily farm and forest land north of Midland. Tests began within 4 h of collection after filtration through coarse paper to remove large solids. The biphenyl compounds were added to the river water without a carrier solvent by evaporating a hexane solution of the material on the inside surface of a glass jar. Rolling the jar after filling with river water gave approximately 100 pg/L solution of the MCBs. These solutions were diluted with river water to approximately 10 and 1pg/L. The TCB was set at 10 and 1pg/L due to ita low water solubility. Aliquots (100 mL) of these river water solutions were transferred to 160-mL serum bottles and sealed with aluminum foil faced Teflon-coated silicone rubber septa. The foil was found necessary to prevent loss of the parent compounds due to volatilization and apparent absorption by the septum. Control samples were treated with 100 mg/L HgClz to enable detection of any abiotic loss of material. All samples were incubated in the dark at 20 "C on a rotary shaker at 100 rpm. Bottles were sacrificed at each time point for analysis. Mineralization of the compounds was observed by trapping 14C02in ethanolamine and 2-methoxyethanol after addition of 0.1 mmol of NaHC03, acidification by the addition of 2.3 mmol of HzS04,and purging with nitrogen. The amount of 14C02trapped was determined by subtracting the amount of 14C-labeled parent compound calculated to have been volatilized from the total radioactivity trapped. The remaining parent compound was determined by extraction of the acidified water solution with methylene chloride followed by concentration and HPLC analysis. The results presented in the figures have been corrected for volatilization of the parent compound. The MCBs were separated from the corresponding benzoic acid metabolites by using a Waters c18 pBondapak reverse-phasecolumn. The solvent system used for elution was composed of 74% methanol, 25% water, and 1% acetic acid at a flow rate of 2 mL/min. Fractions of the HPLC eluent were collected for liquid scintillation counting to determine the parent compound and metabolite concentrations. Aquasol liquid scintillation cocktail (New England Nuclear, Boston, MA) was used throughout this study for 14Cdeterminations. The retention times of the parent compounds under the above conditions were about 6.5,7.3,7.3,5.3, and 13.3 min for 2-MCB, 3-MCB, 4-MCB, BP, and TCB, respectively. The HPLC retention time of the CBA under the above conditions was about 3.5 min, close to the solvent front. Consequently, a Lichrosorb RP-8 column with a solvent system composed of 59.75% (volume) H20, 39.75% methanol, and 0.5% acetic acid was used to isolate CBAs. This allowed a retention time of about 7.5 min for 2-CBA. A 6-min retention time for 4-CBA was observed when the solvent system was composed of 44.75% HzO, 54.75% methanol, and 0.5% acetic acid. Millipore Total Count paddles (Millipore Corp. Bedford, MA) were used to estimate the baderial population of the river water. They are reported to be equivalent to standard plate counts (22) in number of colony forming unit (CFU). Dilutions were prepared by using Millipore dilution kits. The paddles were incubated at 20 "C for 3 days prior to counting. Colonies were counted visually. An average of 6900 CFU/mL in the river water was observed on March 10, 1980. Results Rapid biodegradation of each of the three isomers of MCB and BP was observed in Tittabawassee River water. The disappearance of the parent compounds, appearance of a metabolite(s) in the methylene chloride extract, and 618
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2-MCB, 0.68 pg/L
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Table 1. Times for Biodegradation of 50%of Initial Concentrations of Biphenyl, 2-, 3-, and 4-Monochlorobiphenyl, and 2,2',4,4'-Tetrachlorobiphenyl at 20 "C time, days, at initial concentration, bg/L, of starting compound date 1 10 100
a
biphenyl 5/19/80 1.5 2 2.5 not set 3/10/80 biphenyl 3/10/80 2.5 2 2-MCB 3 3.5 3-MCB 3/10/80 2 3 3/10/80 4-MCB a a 3/10/80 TCB N o degradation observed over 98 days.
3 not set 3.5 4 5 a
the evolution of 14C02are shown in Figures 1-4. The unextracted activity, that which is associated with the cells or very water soluble, is in the raffinate. A comparison of the yields of 14C02from the chlorinated ring of the MCBs and biphenyl at initial concentrations of about lpg/L is shown in Figure 5. The three MCB isomers biodegraded at approximately the same rate at all concentrations, although an apparent lag period was noted at the higher concentrations. Biphenyl was observed to biodegrade somewhat faster than the MCBs. No degradation was observed for TCB, a less than 1% yield of l4COZ.No metabolites were observed by HPLC analysis of the TCB extract after a 98-day incubation. The approximate elapsed times for 50% removal
100 3.MCB, 0.71 pg/L
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of the parent compounds are listed in Table I. A metabolite, believed to be the corresponding chlorobenzoic acid based on the HPLC retention time, was observed from degradation of each of the MCBs. Up to 50% of the initial radioactivity was found as metabolite. The metabolite was observed to degrade more slowly than the parent compound with release of 14C02. No appreciable amount of an extractable metabolite was observed from biphenyl. In addition, a portion of the radioactivity initially added was apparently incorporated in the cell mass or present in some other form which was not extracted by methylene chloride. A material balance was determined for each sample by summing the 14C02trapped, radioactivity extracted, and that remaining in the river water raffinate. The mean accountability of 14Cwas 92.2% with a standard deviation of 12.6 and a range of 58-128%. The mean recovery of parent compound from all control samples was 94.7%. A mean of 5.4% of the radioactivity was apparently volatilized during purging and recovered in the ethanolamine traps in the initial sample workup. This amount of volatilization is approximately what is predicted from the air/water partition coefficients for MCBs, BP, and TCB (23).
Discussion Visual inspection of the results, Figures 1-4, clearly show that the rate of MCB degradation increases with time. The increased rate can be attributed to either an increase in
0
5
10
15 20 Time (Days)
25
30
50
Figure 3. Distribution of radioactivity in biodegradation of 4-MCB: (0) parent 4-MCB, (A)“CO, recovered, (0) metabolite observed, and (A) activity remaining in the river water raffinate.
the population of MCB or BP degrading species or an increase in activity due to increased concentrations of the appropriate enzymes. In this study, the low concentration of added chemical relative to the background TOC in the river water was not expected to cause an appreciable increase in the total bacterial population. All three isomers of MCB were degraded in river water samples at similar rates. The effect of concentration from about 1 to 100 bg/L was greater than the effects of chlorine position on the time for 50% degradation. Previous investigators have examined degradation rates for the MCBs and found essentially the same thing-the degradation rates for the three MCBs vary by only a factor of 2 and are about half that observed for biphenyl (2-4). The similarity of rates of primary degradation for the different compounds is also consistent with the calculations and observations of Kennedy et al. (24) on in vitro metabolism of MCBs by hepatic microsomal cytochrome P-450. They found no differences in the reactivity of the nonchlorinated ring of the different isomers. In addition, extended Huckel molecular orbital calculations by Kennedy et al. (24) indicated the net electronic changes on the unchlorinated ring of the MCBs are close to those calculated for BP. Thus, electrophilic attack on the unchlorinated ring would also be expected to occur at similar rates during bacterial metabolism. The yield of I4CO2from mineralization of the chlorinated ring was similar for all three isomers of MCB. A lag period of about one week was observed before appreciable I4CO2 Envlron. Sci. Technoi., Vol. 17, No. 10, 1983
619
loo I
1
BP. 0.78 pg/L
80
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BP, 1.06pg/L
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Time, Days
Figure 5. Comparison of 14C02 released on biodegradation of approximately 1 pglL solutions of (A)BP, (0)2-MCB, (A)3-MCB, (0) 4-MCB, and ( 0 )TCB.
phase after extraction. BP yielded no appreciable amounts of extractable metabolites. Apparently benzoic acid and its related metabolites were rapidly metabolized to COz and cell mass.
Acknowledgments We thank C. G. Mendoza for excellent analytical advice, W. J. Maier and W. B. Neely for stimulating discussions, and H. T. Bell for assistance in preparing the manuscript.
60 40
Registry No. 2-MCB, 2051-60-7;BP, 92-52-4;TCB, 2437-79-8; 3-MCB, 2051-61-8; 4-MCB, 2051-62-9.
Literature Cited
Time (Days)
Flgure 4. Distribution of radioactivity in biodegradation of BP (0) parent BP, (A)I4CO2recovered, (0) metabolite observed, and (A) activity remaining in river water rafflnate. The set at 0.78 pg/L was concurrent with the MCBs, March 10, 1980. The remaining samples were set May 18, 1980.
was recovered. Apparently, the growth of the microorganisms degrading the chlorinated ring is significantly slower than the growth of microorganisms that degrade the parent MCB. The slow growth rate after a lag period is consistent with literature reports on the biodegradation of chlorobenzoates (19,20).The mineralization rates observed in this study are qualitatively the same as those reported by Boethling and Alexander (12) for 4-chlorobenzoic acid (CCBA). They recovered 48% of theoretical 14C02in 8 days from a stream water sample treated with 4.7 pg/L 4-CBA with slower mineralization at higher concentrations. We observed metabolites that were consistent with the corresponding CBA in all of the experiments with additional carbon-14 remaining in the water 820
Environ. Scl. Technol., Vol. 17, No. IO, 1983
(1) Furukawa, K.; Tonomura, K.; Kamibayashi, A. Appl. Enuiron. Microbiol. 1978, 35, 223-227. (2) Ahmed, M.; Focht, D. Can. J . Microbiol. 1973,19,47-52. (3) Wong, P. T. S.; Kaiser, K. L. E. Bull. Enuiron. Contam. Toxicol. 1975, 13, 249-255. (4) Reichardt, P. B.; Chadwick, B. L.; Cole, M. A.; Robertson, B. R.; Button, D. K. Enuiron. Sci. Technol. 1981,15,75-79. (5) Shiaris, M. P.; Sayler, G. S. Enuiron. Sci. Technol. 1982, 16, 367-369. (6) Clark, R. R.; Chian, E. S. K.; Griffin, R. A. App. Enuiron. Microbiol. 1979, 37, 680-685. (7) Branson, D. R., The Dow Chemical Co., Midland, MI, personal communication, 1979. (8) Tucker, E. S.; Saeger, V. W.; Hicks, 0. Bull. Enuiron. Contam. Tox. 1975,14,705-713. (9) Baxter, R. A.; Gilbert, P. E.; Lidgett, R. A,; Mainprize, J. H.; Vodden, H. A. Sci. Total Enuiron. 1975, 4 , 53-61. (10) Hites, R. A. J . Chromatogr. Sci. 1973, 11, 570-574. (11) Ballschmiter, K.; Unglert, Ch.; Neu, H. J. Chemosphere 1977,6,51-56. (12) Boethling, R. S.; Alexander, M. Appl. Enuiron. Microbiol. 1979,37, 1211-1216. (13) Haller, H. D. J . Water Pollut. Control Fed. 1978, 49, 2771-2777. (14) Reineke, W.; Knackmuss, H.-J. Biochim. Biophys. Acta . . 1978,542,412. (15) DiGeronimo, M. J.; Nikaido, M.; Alexander, M. Appl. Enuiron. Microbiol. 1979. 37. 619-625. (16) Knackmuss, H.-J.; Reineke,' W. Chemosphere 1973, 2, 225-230. (17) Spokes,J. R.; Walker, N. Arch. Microbiol. 1974,96,125-134. (18) Hartmann, J.; Reineke, W.; Knackmuss, H.-J. Appl. Enuiron. Microbiol. 1979, 37, 421-428. (19) Ruisinger, S.; Klages, U.; Lingens, F. Arch. Microbiol. 1976, 110, 253-256. (20) Haller, H. D.; Finn, R. K. Eur. J . Appl. Microbiol. Biotechnol. 1979,8, 191-205.
Envlron. Sci. Technol. 1983, 17, 621-624
(21) Shamat, N.; Maier, W. J. J . Water Pollut. Control Fed. 1980,52, 2158. (22) Cotton, R. A.; Sladek, K. J.; Sohn, B. I., World Congress of Environmental Medicine and Bioloy, Paris,France, July 1974. (23) Bailey, R. E.; Rhinehart, W. L.; Gonsior, S. J.; Batchelder, T. L.; Mendoza, C. G.; Neely, W. B., Second Annual Meeting
of the Society for Environmental Toxicology and Chemistry, Arlington, VA, Nov 1981. (24) Kennedy, M. W.; Carpentier, N. K.; Dymerski, P. P.; Adams, S. M.; Kaminsky, L. S. Biochem. Pharmacol. 1980, 29, 727-736. Received for review February 7,1983. Accepted June 15,1983.
NOTES Is the Direct Mutagenic Activlty of Diesel Particulate Matter a Sampling Artifact? Terence H. Rlsby" Division of Environmental Chemlstry, Department of Envlronmental Health Sciences, The Johns Hopkins University School of Hyglene and Public Health, Baltimore, Maryland 21205
Samuel S. Lestz Department of Mechanical Engineering, The Pennsylvania State University, University Park, Pennsylvania 16802 Some of the direct microbial mutagenic activity of diesel particulate matter has been assigned to the presence of nitro derivatives of polynuclear aromatic hydrocarbons. A number of researchers have suggested that these compounds may be produced during sample collection. This paper discusses their formation on the basis of simple chemical kinetic and collision phenomena.
Introduction Many reports have recently appeared in the literature that demonstrate, by in vitro microbial assays, the direct mutagenic activity of extracts of diesel particulate matter: examples of this work are contained in three books, published during the past two years (1-3). The observation that these samples do not require mammalian enzymes for activation of mutagenic activity was interesting since engine combustion sources are known to produce polynuclear aromatic hydrocarbons (PAHs) that require microsomal activation (4). It was therefore apparent that the diesel sorbate contains additional classes of mutagens. One such class of direct mutagens is nitroaromatics, and some of these compounds have been identified on diesel particles [1-nitropyrene, nitromethylanthracene, hydroxynitrofluorene, nitrodehydropyrene, and 2-nitrofluorene (5)]. These rapid predictive microbial assays have been confirmed with mammalian cell culture assays, skin painting, and intratracheal instillations (1-3). However, these results have not been confirmed by studies in which various types of laboratory animals were exposed to diluted total diesel exhaust; no increased incidence of carcinomas was demonstrated (1-3). There was morphological and biochemical evidence of lung tissue injury and adaption or repair after inhalation of very high levels of diesel exhaust (6). While this difference between the results obtained with in vitro and in vivo assays and with animal exposure studies is not without precedent, it has served to cloud the understanding of the potential health effects of diesel exhaust. There are many possible explanations for these differences in results such as relative bioavailability of the adsorbates on the diesel particle (7), detoxification mecha0013-936X/83/0917-0621$01.50/0
nisms (3), number of animals exposed (3),lack of typical atmospheric interactions (2),and long-term retention of particles (2). However, a simpler explanation may be that there is a chemical difference between the agents of exposure in in vitro and in vivo assays and in inhalation exposure. This chemical difference may arise during sampling, and if so, the direct mutagenic activity of diesel particles is therefore an artifact. Theoretically it should be possible to investigate whether sampling artifactual processes are causing the problem by performing exposure studies using reaerosolized diesel particles; in practice these studies have not proved successful. Diesel particles have a mean particle diameter of 0.027 pm (81, and the mean diameter of reaerosolized diesel particles in the only study reporting such exposure was 2.8 pm (9). It is therefore difficult to use such data to support conclusive argumenta for or against the premise that the activity is produced as a sampling artifact. Collection of diesel particles by different sampling methods (e.g., electrostatic precipitation) represents an alternative approach. The direct microbial mutagenic activity of the extract from e!wtrostatically precipitated diesel particles was found to be not significantly different from the direct activity of filtered particles (10). These results suggest that the direct microbial mutagenic activity is not due to sampling artifactual reactions. Alternatively, the reduction in direct mutagenic activity from sampling artifactual reactions could be compensated by reactions with ozone: ozone is produced by the corona discharge in electrostatic precipitators. Pitts et al. have shown that both ozone and nitric acid react with polynuclear aromatic hydrocarbons to produce strong direct microbial mutagens (11,12). The direct microbial activities of an oxide and a nitro derivative of benzo[a]pyrene have been shown to be comparable [l-or 3-nitrobenzo[a]pyrene, 3600-5300 revertants/pg (11, 13); benzo[a]pyrene 4 5 oxide, 1600 revertants/pg (12)].In addition ozone will rapidly oxidize nitric oxide to nitrogen dioxide, which has been postulated as the reactant, in the presence of small amounts of water vapor, for the formation of direct mutagens (nitropolynuclear aromatic compounds). Therefore the direct microbial mutagenic activities of electrostatically
0 1983 American Chemical Society
Environ. Sci. Technol., Vol. 17, No. 10, 1983 621
