Environ. Scl. Technol. 1986, 20, 1002-1007
Evans, W. C. Nature (London) 1977,270, 17-22. Healy, J. B., Jr.; Young, L. Y.; Reinhard, M. Appl. Environ. Microbiol. 1980, 39, 436-444. Aftring, R. P.; Chalker, B. E.; Taylor, B. F. Appl. Environ. Microbiol. 1981, 41, 1177-1183. GrbiE-GaliE, D. A p p l . Environ. Microbiol. 1983, 46, 1442-1446. Braun, K.; Gibson, D. T. Appl. Environ. Microbiol. 1984, 48, 102-107. GrbiE-GaliE,D.; Young, L. Y. Appl. Environ. Microbiol. 1985, 50, 292-297. Kuhn, E. P.; Colberg, P. J.; Schnoor, J. L.; Wanner, 0.; Zehnder, A. J. B.; Schwarzenbach, R. P. Environ. Sei. Technol. 1985, 19, 961-968. Wilson, J. T.; McNabb, J. F. E O S , Trans. Am. Geophys. Union 1983, 64, 505-506. Jamison, V . W.;Raymond, R. L.; Hudson, J. O., Jr. In Proceedings, Third International Biodegradation S y m posium; Sharpley, J. M.; Kaplan, A. M., Eds.; Applied
Science Publishers: Essex, U.K., 1976; pp 187-196. (30) Lee, M. D.; Ward, C. H. In Proceedings, Hazardous Material Spills Conference: Prevention, Behavior, Control and Cleanup of Spills and Waste Sites; Nashville, TN, 1984; pp 98-103. Received for review November 11,1985. Accepted M a y 1,1986. This study was supported by the United States Air Force through Interagency Agreement R W57930615-01-1 with the U.S. Environmental Protection Agency. T h e work was funded under Cooperative Agreement CR-811146 between the R. S. Kerr Environmental Research Laboratory and the University of Oklahoma at Norman. Although the research described in this article has been supported in part by the US.Environmental Protection Agency under assistance agreement number CR-811146 t o the University of Oklahoma and under in-house programs, it has not been subjected to the Agency's peer and administrative review and, therefore, does not necessarily reflect the views of the Agency, and no official endorsement should be inferred.
Degradation of Phenol and Chlorophenols by Sunlight and Microbes in Estuarine Water Huey-Min Hwang" and R. E. Hodson
Department of Microbiology and Institute of Ecology, University of Georgia, Athens, Georgia 30602 R. F. Lee
Skidaway Institute of Oceanography, Savannah, Georgia 314 16 The rates of photolysis and microbial degradation of phenol, p-chlorophenol, 2,4-dichlorophenol, 2,4,5-trichlorophenol, and pentachlorophenol in estuarine water were determined. Photolysis was the primary transformation process for the polychlorinated phenols with photolysis rate constants in surface estuarine water ranging from 0.3 to 1.2 h-l and half-lives ranging from 0.6 to 3 h. Dichlorophenol photolysis rates were 20430% higher in estuarine water than in distilled water, indicating a photosensitized reaction. There was no microbial (dark) degradation of polychlorinated phenols during short incubation periods (up to 3 days). The photoproducts of polychlorinated phenols were rapidly degraded by microbes. Microbial degradation was the primary process for transformation of phenol and p-chlorophenol. In the summer the microbial and photolysis transformation rate constants for phenol were 0.03 (tllz= 28 h) and 0.016 h-l (tllz= 43 h), respectively. Winter photolysis and microbial degradation rates were lower than the summer values. ~~
Introduction As a result of man's agricultural and industrial activities, phenol and various chlorophenols are found in estuaries near urban areas (1). The fate of these compounds is of interest because of their toxicity to marine life and their threat to human health (2). Studies of the fate of chlorophenols in outside enclosures, so-called microcosms or mesocosms, have shown that microbial activity and photolysis are the processes primarily responsible for degradation of chlorophenols in aquatic environments (3-5). Microbial metabolism of phenol and chlorophenols has been studied in both freshwater and marine environments (3,644. The degradation followed first-order kinetics after an initial lag (6). Repeated additions of chlorophenols to marine or freshwater mesocosms resulted in an increase in their degradation rates, suggesting an increase in the 1002
Environ. Sci. Technol., Vol. 20, No. 10, 1986
population of chlorophenol-degrading microbes (3, 4). Addition of more chlorines to the chlorophenol ring resulted in slower biodegradation rates (6). Microbial metabolism of chlorophenols in aerobic waters involves oxidative dechlorination and hydroxylation. The metabolism of pentachlorophenol produces trichlorophenol, while chlorocatechols are intermediates in the metabolism of monochlorophenols (1,9). Ring cleavage can also occur with COz as the final product. Photolysis studies have been carried out on a number of chlorophenols. Photolysis of pentachlorophenol in distilled water resulted in a number of degradation products including tetrachlorophenols, trichlorophenols, chlorinated dihydroxybenzenes, and nonaromatic fragments such as dichloromaleic acid (10). Photolysis of monochlorophenols and dichlorophenols resulted in dechlorination and formation of catechol and other hydroxy benzenes (11, 12). In natural waters direct photolysis can be slower than in distilled water due to attenuation of light by dissolved substances and suspended particulates (13). The beam attenuation coefficient and diffuse attenuation coefficient were found to be related to light attenuation by dissolved substances and suspended particulates, respectively (13). For some compounds photolysis rate is higher in natural waters than in distilled water due to the presence ,of photosensitizers, e.g., humics, which results in indirect photoreactions (14). Both sunlight and microbial activity change during the year. For example, surface irradiance is reduced in the winter relative to summer so benzo[a]pyrene photolysis rates are lower in the winter (15). Microbial degradation rates of a variety of compounds have been demonstrated to be lower in the winter, presumably due to the lower temperatures. Thus, during the year there can be changes in the relative importance of photolysis and microbial degradation as they affect the fate of xenobiotics. In our
0013-936X/86/0920-1002$0 1.50/0
0 1986 American Chemical Society
studies the relative importance of microbial and photochemical degradation of phenol and a series of chlorophenols in estuarine water was determined. Changes in microbial degradation and photolysis rates were related to changes in biological, chemical, and physical properties of the water. Experimental Section Surface water samples were collected during high-tide periods from Skidaway River, an estuarine river located near Savannah, GA. Water -samples were collected in acid-washed, polyethylene containers. Assays were initiated within 1 / 2 h of collection. At time of sampling, temperature, pH, and salinity were measured. Chemicals. Ring-ULJ4C-labeled p-chlorophenol(11.61 mCi/mmol), 2,4-dichlorophenol (10.7 mCi/mmol), and 2,4,5-trichlorophenol(0.80mCi/mmol) were obtained from Pathfinder Laboratories Inc. Ring-UL-14C-labeledphenol (58 mCi/mmol) and pentachlorophenol (8.8 mCi/mmol) were obtained from California Bionuclear Corp. Valerophenone and other unlabeled chemicals were obtained from Aldrich Chemical Co. Malachite green leucocyanide actinometer (MGLC) was obtained as a gift from Dr. R. G . Zepp of the U. S. Environmental Protection Agency Laboratory in Athens. 3H-LabeledD-glucose (30 Ci/mmol) and 14C-labeledsodium bicarbonate (50 mCi/mmol) were obtained from New England Nuclear Corp. Incubation and Degradation Measurements. Radiolabeled compounds dissolved in acetone were added to 60 mL of water sample in 150-mL quartz Erlenmeyer flasks (Quartz Scientific, Inc.). Acetone was a t a concentration of 1 X M, and at this concentration there was no evidence of photosensitization by acetone in distilled water of the compounds. These flasks allowed more than 85% transmission of light of wavelength greater than 260 nm. Approximately 0.1 pCi of the selected radioactive compound was added to each flask. Final concentration of each compound in the flasks was adjusted to 25 pg L-l by the addition of unlabeled compound. Flasks were stoppered (silicone; VWR Scientific Inc.) and suspended in an outdoor tank through which estuarine water was continuously circulated, and the water level in the flask was 3 cm below the surface. Flasks were covered with aluminum foil for dark experiments. Formaldehyde (0.37% final concentration) was added to distilled and estuarine water in sunlight for photolysis only experiments. Ultraviolet absorption by the formaldehyde solution at this concentration was found to be negligible. Parent compound disappearance (transformation) and 14C02appearance (mineralization) were determined for each experiment. Since phenol and p-chlorophenol photolyzed slowly, they were exposed to sunlight for up to 3 days; transformation studies of other chlorophenols were conducted under midday sunlight (10 a.m.4 p.m.; eastern standard time). Mineralization studies used water samples incubated for 24, 48, or 72 h to allow production of sufficient 14C02to detect. The 14C02produced was determined as described elsewhere (6,16). To determine disappearance of parent compound, samples were taken to pH 2 with 4 N HzSO4, followed by extraction of parent compounds and degradation products by two extractions with ethyl acetate. Extraction efficiency was greater than 95% for all compounds. Extracts were concentrated by evaporation under nitrogen and applied to silica gel thinlayer chromatography plates (E. Merck) and run with a solvent system of hexane-acetone (1:l v/v). There was some minimal, unavoidable loss of chlorophenol extracts (611%) during the concentration step. Parent compounds and degradation products were scraped from thin-layer
plates, and their radioactivity was determined with a liquid scintillation counter (Packard TRI-carb; 300C). Analyses of malachite green leucocyanide used a Bausch & Lomb Model Spectronic 20 spectrophotometer, following the procedures described by Miller and Zepp (13). The screening factor (defined later in this section) was determined by exposing the valerophenone solutions (0.01 mM) to midday sunlight for up to 1 h. Analyses for valerophenone were carried out with high-performance liquid chromatography (Micromeritics Model 7000B) using a reverse-phase column (10 pm ODS/Spherisorb Excalibar Inc.), mobile phase of 70:30 methanol/water, flow rate of 2.0 mL mi&, and ultraviolet-visible detector (wavelength 260 nm). Each radiolabeled compound solution was incubated in triplicate; valerophenone solutions in duplicate were incubated simultaneously on different days under nearly identical conditions of irradiance and temperature. Each experiment was repeated a t least once, and the mean values are presented under Results. Transformation (disappearance of parent compound) rate constants and mineralization (14C02production) rate constants were calculated by assuming the reaction were first order (6,17). The first-order rate expression is given by In (Co/C) = k,t
(1)
where C, and C refer to the concentrations of the measured compound at to and t , respectively, and k, is the first-order photolysis rate constants, in units of h-l or day-l. The half-lives of the compounds were determined by til2 = 0.693/kp
(2)
The first-order rate constants were corrected for abiotic loss in darkness (less than 5% of the total rate) and light attenuation by dissolved organic material. Experiments were conducted on sunny days. Solar radiation was integrated hourly with a radiometer (LI-COR Inc., Model LI-550B; active range 400-700 nm). The light screening factor (S)was calculated as the ratio of the valerophenone photolysis rate constant in estuarine water to the rate constant in distilled water (18). Properties of Estuarine Water. Concentrations of suspended particulates in estuarine water were determined by filtering 1L of estuarine water through predried and preweighed glass-fiber filters (GF/C, Whatman). After filtration the filters were dried and reweighed. Particulate-free estuarine water was obtained by ultracentrifugation of a water sample at lOOOOOg (Beckman Ultracentrifuge Model L5-40) for 1 h. The particulate-free estuarine water was used in screening factor determinations. Absorption spectra of the particulate-free estuarine water were obtained with a Beckman spectrophotometer, Model DU-6. Diffuse attenuation coefficients were determined by using a malachite green leucocyanide actinometer as described by Miller and Zepp (13). Flasks containing malachite green leucocyanide (2 X M) in estuarine water were suspended a t different depths in the estuary. The amount of malachite green formed a t each depth after a 10-min incubation period was determined with a spectrophotometer (622 nm). The diffuse attenuation coefficient was computed by the equation discussed by Miller and Zepp (13):
K = In
[(A622
at Zl)/(A622 a t z2)l(z2 - 2,)
(3)
where A,&) was the absorbance of the actinometer at 622 nm after exposure at depth 2. The distribution factor (D)is the mean path length of light in an horizontal layer of the water sample divided by thickness of the layer (13), Envlron. Scl. Technol., Vol. 20, No. 10, 1986
1003
and in our experiments it was 1.1. For analysis of particulate organic carbon and particulate nitrogen, 500 mL of water was passed sequentially through two Gelman type A glass-fiber filters which had been precombusted a t 500 "C. The filters were rinsed with 0.01 N HC1 to remove inorganic carbon and dried and stored in a 60 "C oven. Particulate organic carbon on the filters was determined by the methods of Menzel and Vaccaro (19). Particulate organic nitrogen was determined by the classical micro-Dumas method as described by Strickland and Parsons (20). The analysis of dissolved organic carbon consisted of the wet oxidation of filtered estuarine water by potassium persulfate in a sealed glass ampule. Samples were subsequently analyzed by the methods of Menzel and Vaccaro (19). Nitrate and phosphate were analyzed by methods described by Strickland and Parsons (20).
Microbial Biomass and Activity Measurements. Microbial heterotrophic activity was determined by adding tracer quantities of tritiated D-glucose (0.45 nmol of D[6-3H(N)]glucose;30 Ci/mmol) to 60 mL of estuarine water in quartz flasks, incubating for various periods up to 1hr, and thGn filtering the sample water on 0.2-pm-pore Gelman filters. Filters were washed with 20 mL of filtered estuarine water, and the filters were counted with a liquid scintillation counter. Similarly the uptake of I4CO2by algae under sunlight was used as an index of autotrophic microbial activity and was determined by adding 1 pCi of sodium [14C]bicarbonate to 60 mL of estuarine water. After 2 h of incubation, the water was filtered on 0.2-pm-pore Gelman filter, and the filter was washed, acidified with concentrated HC1, and radioassayed as above. Turnover times of glucose and bicarbonate in estuarine water were calculated as t / f where t is the incubation time and f is the fraction of added label assimilated in incubation time t , and the turnover times served as an index of metabolic activity. A correction factor of 0.5 for [3H]-~-glucose respiration to 3H20was applied for final calculation of turnover time (21). Total microbial biomass was estimated by measurements of particulate adenosine triphosphate (ATP) (22). Bacterial numbers in water samples were determined by the acridine orange direct count (AODC) microscopic method (23). Chlorophyll a concentrations, used as a measure of phytoplankton biomass, were determined fluorometrically by using the method of Yentsch and Menzel (24) as described in Strickland and Parsons (20). Results Some biological, chemical, and physical properties of the Skidaway River are listed in Table I. The pH, absorbance of the particulate-free estuarine water, particulate organic carbon concentration, and dissolved organic carbon concentration remained fairly constant throughout the year. Temperature, surface irradiance, and diffuse attenuation coefficient changed markedly during the year. The microbial transformation and mineralization rates (dark), phototransformation and photomineralization rates (poisoned, light), and simultaneous microbial degradation and photolysis rates (light) are given for each compound in Table 11. Photolysis rates were determined in both distilled and estuarine water. No degradation was observed in poisoned dark samples. The relative rates of photolysis decreased in the order 2,4,5-trichloropheno1, 2,4-dichlorophenol, pentachlorophenol, p-chlorophenol, and phenol (Table I1 and Figure 1). The phototransformation of the compounds was first order with respect to concentration (Figure 1). 1004
Environ. Sci. Technol., Vol. 20, No. IO, 1986
-1
4 1
oentachloroohenoi I 2 4-dtchlorophenol ~1,24.5-tri~hlorophenol n
1
0
; ; ' ; ' i ' i
O'
Time (hours1 Figure 1. Photolysis of chlorophenols. Chlorophenols were added to the estuarine water at a concentration of 25 pg L-' in December. Temperature was 14 OC, and pH was 7.6. Phototransformation equations were In c = 3.22 -0.27t (pentachlorophenol), In c = 3.22 - 0.44t (2,4-dichlorophenol), and In c = 3.22 - 0.65 t (2,4,5-trichlorophenol), where c is the concentration of the chlorophenol at time t . Vertical bars represent 1 standard deviation with n = 3.
* Distilled water a 0
Poisoned estuarine water Unpoisoned estuarine water
1
2
3
4
5
Time (hours1 Figure 2. Degradation of 2,4-dichlorophenol. Dichlorophenol was added to the water samples at a final concentration of 25 pg L-' in December. Temperature was 14 OC, and pH was 7.6. Transformation equations were In c = 3.22 - 0.21t (photolysis in distilled water), In c = 3.22 - 0.38t (photolysis in poisoned estuarine water), and In c = 3.22 - 0.44t (photolysis and microbial degradation in unpolsoned estuarine water), where G is the concentration of dichlorophenol at time t . Vertical bars represent 1 standard deviation with n = 3.
The phototransformation rate constants for 2,4-dichlorophenol, 2,4,5-trichlorophenoI,and pentachlorophenol ranged from 0.3 to 1.2 h-l with half-lives ranging from 0.6 to 3 h. Photolysis of phenol and p-chlorophenol was slower, and half-lives ranged from 43 to 118 h. Similar differences were observed for photomineralization with half-lives ranging from 6 to 14 days for dichlorophenol, trichlorophenol, and pentachlorophenoland from 16 to 334 days for phenol and p-chlorophenol. Phototransformation and photomineralization rates for all compounds were higher in summer than in the winter (Table 11). For example, the phototransformation rate constant of 2,4,5-trichlorophenolincreased from 0.65 to 1.2 h-' from winter to summer. The higher photolysis rate of dichlorophenol in estuarine water relative to distilled water suggested a photosensitized reaction (Figure 2). The lower photolysis rate of pentachlorophenol in estuarine water was presumably due to chloride inhibition of photolysis (25). Photolysis rates for phenol, chlorophenol, and trichlorophenol were the same in distilled and estuarine water when corrected for attenuation of light, i.e., screening factor.
Table I. Physical, Chemical, and Biological Properties of Skidaway River Water: 1983-1985 chemical Droperties
physical properties PH temperature, "C suspended particulates, mg L-' K330,cm-lb Zoo.cmc
7.7 (7.4-8.0) 21.7 f 6.3 (7-29) 31.3 f 17.1 (12.9-68.3) 0.22 (Oct 1983), 0.05 (Feb 1984) 21 (Oct 1983), 100 (Feb 1984) 0.15 (Oct 1983), 0.01 (Feb 1984) 0.05 f 0.02 (0.030-0.096) 6.5 x (Oct 1983), 0.6 X (Feb 1984) 0.85 f 0.04 (0.82-0.88)
dissolved organic carbon, mg/L 4.8 f 0.3 particulate organic carbon, mg/L 0.9 f 0.1 particulate organic nitrogen, mg/L 0.20 f 0.05 4.6 f 1.3 C/N 0.7 f 0.3 phosphate, Fg-atom/L nitrate, pg-atom/L 1.4 f 0.3 22.8 f 2.4 (18-25) salinity, %OB
Biological Properties
season and temp, "C
acridine orange direct count (AODC), (number/mL) x 106
particulate ATP, pLg/L
chlorophyll a, Fg/L
summer, 28.5 winter, 13.7
7.0 f 1.4 (5.5-8.3) 4.5 f 2.9 (1.1-9.8)
1.6 f 0.6 (1.0-2.3) 3.0 f 1.1 (1.8-4.6)
7.9 f 2.2 (5.5-9.8) 8.3 f 4.2 (4.2-12.8)
[I4C]bicarbonate uptake turnover specific turnover rateh time, h 289 f 220 (133-444) 1370 f 923 (573-2312)
4.4 0.9
[3H]glucoseuptake specific turnover time, h turnover rate' 2.1 f 0.6 (1.6-2.6) 3.7 f 0.7 (3.0-4.6)
7.1 6.0
aExpressed as the mean f standard deviation (range). bDiffuse attenuation coefficient at 330 nm. cZgg= 4.6/X, photic zone depth. The subscript 99 means the depth of 99% attenuation of surface irradiance. dComponent of K due to light attenuation by suspended particulates. a330 is the absorbance (l-cm path length) of the particulate-free estuarine water. D is the distribution coefficient in estuarine water. e Specific attenuation coefficient due to suspended particulates. f Screening factor is the ratio of valerophenone photolysis rate in estuarine water to that in distilled water. gym, parts per thousand. hDefined as the turnover rate divided by chlorophyll a concentration (h-' L pg-I X 'Defined as the turnover rate divided bv AODC numbers (h-' mL cell-' X 10-9.
The primary process responsible for the transformation and mineralization of phenol and p-chlorophenol during summer months was microbial degradation (Table 11). Moreover, transformation and mineralization of these compounds were slower in the light than in the dark. Thus, the half-lives for p-chlorophenol in the dark and light were 11 and 28 h, respectively. This result suggests inhibition of microbial activity by sunlight in surface waters, possibly due to the inhibition by solar radiation (26)or toxicity of photoproducts (27). However, our data (not shown) indicated that neither was the case. Since in the winter microbial degradation rates of phenol and chlorophenol were the same in the light and dark (Table 11), we speculate that the higher concentrations of algal photosynthetic products in the summer may have inhibited biodegradation of phenol and chlorophenol. Evidence that an algal photosynthetic product inhibited biodegradation of chloroaniline photoproducts in a freshwater lake was noted in an earlier study (28). In the time periods used (up to 3 days) there was no significant microbial, i.e., dark degradation, of dichlorophenol, trichlorophenol, or pentachlorophenol. Microbial degradation rates of phenol and p-chlorophenol were low in the winter when temperatures were 10-14 "C. The dark transformation rate constants for p-chlorophenol in summer and winter were 0.06 and 0.006 h-l, respectively. As bacterial numbers were not significantly different between winter and summer (Table I), it is assumed that seasonal differences in microbial degradation rates were due to temperature changes. Incubation of compounds in sunlit estuarine water allowed microbial degradation and photolysis to act simultaneously. After photolysis of polychlorinated phenols, there was microbial as well as abiotic mineralization of the photoproducts. The mineralization half-life of pentachlorophenol in sunlit estuarine water was 3 and 6 days in unpoisoned and poisoned samples, respectively (Table 11). The winter transformation half-lives in the light and dark of p-chlorophenol were 63 and 116 h, respectively
(Table 11), showing that photolysis was more important than microbial degradation in the winter. For the polychlorinated phenols there was a pronounced decrease in the microbial degradation of the photoproducts in the winter. For p-chlorophenol, microbial degradation was the major degradative process in the summer, while photolysis contributed to degradation in the winter. The lower transformation rate for p-chlorophenol in the winter relative to summer was due to a decrease in both microbial activity (Table I) and surface irradiance (Table 11). Surface irradiance was 2.7 and 5.9 einsteins m-2 h-l (wavelength 400-700 nm) in the winter and summer, respectively, while p-chlorophenoltransformation rates were 0.01 and 0.03 h-l, respectively (Table 11). For phenol, microbial degradation was the primary removal process in both winter and summer.
Discussion Photolysis rates were high for all polychlorinated phenols (2,4-dichlorophenol, 2,4,5-trichlorophenol, and pentachlorophenol) and low for phenol and p-chlorophenol in estuarine or distilled water. Other investigators have reported high photolysis rates for polychlorinated phenols in natural waters. For example, pentachlorophenol at a depth of 3.8 cm in a freshwater stream or in surface seawater had a photolysis rate constant of 0.29 h-' (tl/z = 2.4 h) ( 4 , 2 5 ) . We determined a k, of 0.37 h-l (tllz= 2 h) a t a depth of 3.0 cm for pentachlorophenol in estuarine water (Table 11). Although photolysis rates of polychlorinated phenols are very high in surface waters, if the entire water column is considered, photolysis rates are much lower. The half-life of pentachlorophenol in a l-m deep freshwater pond was 1.5-3 days (29), while in a 5.5-m deep marine mesocosm the half-life was 22 days (5). The calculated half-life of pentachlorophenol a t a depth of 40 cm in a freshwater stream was 625 days (29). The long half-life was due to attenuation of light by dissolved and particulate substances (4). Similar calculations using the diffuse attenuation coefficient of our estuary showed that photolysis of polyEnviron. Sci. Technol., Vol. 20, No. 10, 1986
1005
Table 11. Photolysis and Microbial Degradation of Phenol and Chlorophenols" compound (water) phenol distilledb estuary (poisoned) estuary estuary (dark) p-chlorophenol distilled estuary (poisoned) estuary estuary (dark) 2,4-dichlor~phenol~ distilled estuary (poisoned) estuary 2,4,5-trichlorophenol distilled estuary (poisoned) estuary pentachlorophenol distilled estuary (poisoned) estuary
season (temp, "C)
midday surface irradiance, E m-2 h-'
transformation rate constant, half-life, h-l h
mineralization rate constant, half-life, day-' day
summer (24) winter (10) summer (24) winter (10) summer (24) winter (10) summer (24) winter (10)
4.9 f 1.3 2.9 f 1.1 4.9 =k 1.3 2.9 f 1.1 4.9 f 1.3 2.9 f 1.1
0.015 f 0.006 0.0040 f 0.0002 0.016 f 0.006 0.006 f 0.001 0.018 f 0.004 0.0074 f 0.0006 0.03 f 0.01 0.011 f 0.004
46 173 43 118 39 94 28 62
0.04 f 0.02 0.0041 f 0.0005 0.04 f 0.02 0.0063 f 0.0008 0.095 f 0.019 0.010 f 0.001 0.4 f 0.2 0.0051 f 0.0008
summer (25) winter (14) summer (25) winter (14) summer (25) winter (14) summer (25) winter (14)
5.9 f 0.6 2.7 f 1.3 5.9 0.6 2.7 f 1.3 5.9 f 0.6 2.7 f 1.3
0.011 f 0.006 0.007 f 0.002 0.015 f 0.007 0.011 f 0.004 0.03 f 0.01 0.011 f 0.004 0.06 f 0.03 0.006 f 0.001
63 99 46 63 28 63 11 116
0.012 f 0.003 0.003 f 0.002 0.013 f 0.004 0.0021 f 0.0005 0.07 f 0.01 0.007 f 0.002 0.293 f 0.009 0.003 f 0.002
summer (25) winter (11) summer (25) winter (11) summer (25) winter (11)
5.7 f 0.2 1.8 f 1.0 5.7 f 0.2 1.8 f 1.0 5.7 f 0.2 1.8 f 1.0
0.82 f 0.06 0.21 f 0.05 1.00 f 0.06 0.38 f 0.02 1.16 f 0.07 0.44 f 0.06
0.8 3 0.7 2 0.6 2
0.09 i 0.03 0.049 f 0.005 0.12 f 0.05 0.05 f 0.01 0.20 f 0.04 0.04 f 0.01
14 6 14 4 17
summer (25) winter (18) summer (25) winter (18) summer (25) winter (18)
5.3 f 0.5 2.3 f 1.2 5.3 f 0.5 2.3 f 1.2 5.3 f 0.5 2.3 f 1.2
1.3 f 0.2 0.61 f 0.08 1.2 f 0.3 0.65 f 0.06 1.4 f 0.1 0.65 f 0.06
0.5 1 0.6 0.5 1
0.10 f 0.03 0.050 f 0.009 0.12 f 0.04 0.05 f 0.01 0.28 f 0.04 0.09 f 0.01
7 14 6 14 3 8
summer (25) winter (11) summer (25) winter (11) summer (25) winter (11)
5.2 f 1.5 1.9 f 0.8 5.2 f 1.5 1.9 f 0.8 5.2 f 1.5 1.9 f 0.8
0.6 f 0.1 0.37 f 0.06 0.37 f 0.08 0.27 f 0.04 0.37 f 0.07 0.27 f 0.04
1 2 2 3 2 3
0.11 f 0.04 0.049 f 0.008 0.12 f 0.05 0.07 f 0.01 0.25 f 0.01 0.10 f 0.02
6 14 6 10 3 7
*
1
16 169 16 110
7 73 2 136 58 224 53 334 10
95 2 231 8
'*C-Labeled compounds were incubated in quartz flasks at a concentration of 25 pg L-l. The flasks with distilled or estuarine water were exposed during days when there was full sunlight, Le., no clouds. Distilled or poisoned estuarine water was treated with formaldehyde (0.4%). Values are expressed as the mean f standard deviation (n = 3). 2,4-Dichlorophenol, 2,4,5-trichlorophenol, and pentachlorophenol were exposed to midday sunlight for 4 h, and half-lives were reported as "light hours". Because of slower photolysis rates, phenol and p-chlorophenol were exposed to sunlight and darkness for up to 3 days. The estuarine rate constants had been corrected for the screening factor. bDistilled water was buffered at pH 7.7 f 0.2 (0.016 M phosphate). CDegradationsof dichlorophenol, trichlorophenol, and pentachloroDheno1in darkness were negligible. (I
chlorinated phenols would be very low at deeper depths (below 20 cm). Thus, for deeper waters degradative processes other than photolysis must be considered. In our short incubations (3 days or less) there was no significant microbial degradation of the polychlorinated phenols, although there was rapid microbial degradation of photoproducts. Longer incubation periods would have presumably shown biological degradation. For example, Pignatello et al. ( 4 ) found biological degradation of pentachlorophenol (in the dark) began 22 days after addition to a freshwater stream. Entrance of polychlorinated phenols into sediments results in enhanced microbial degradation of the compounds. Degradation rates are higher in the sediment than in the water due to higher microbial activity in the sediment (6, 30). The dark mineralization half-life of trichlorophenol added to a sediment-water slurry from the same estuarine river used in the present study was 23 days (6). Phenol and p-chlorophenol showed slow photolysis rates but high microbial degradation rates when compared with polychlorinated phenols. Exposure of monochlorophenols in water to ultraviolet light produces various photopro1006
Environ. Sci. Technol., Vol. 20, No. 10, 1986
ducts, including catechol and other hydroxybenzenes (11). However, in our estuarine water microbial degradation, particularly in the summer, accounted for most of the transformation of phenol and p-chlorophenol. The microbially mediated rates for both transformation and mineralization of phenol and p-chlorophenol were lower in winter than in the summer. Lower winter turnover rates have been found for several organic substrates in estuarine water (31). Although the abundance of chlorophenol degraders in our studies were not determined, other work has shown that the proportion of degraders of xenobiotic compounds to total bacteria is relatively small (32). As microbial numbers were comparable in the two seasons while microbial turnover of glucose was faster in summer than in the winter (Table I), we speculate that temperature was affecting microbial degradation rates of chlorophenols. The concentration of suspended particulates was higher in the summer, resulting in a summer diffuse attenuation coefficient (at 330 nm) 4 times higher than the winter value. Since photolysis rates for chlorophenols were higher in the summer, it appears that the increase in irradiance
during the summer was a more important factor than the increased light attenuation in determining the photolysis rates in surface water. For deeper waters the increased light attenuation by suspended particulates in the summer would be a major factor affecting photolysis rates. 2,4-Dichlorophenol had a higher photolysis rate in estuarine water than in distilled water, indicating a photosensitized reaction. It has been well documented that humic substances can be responsible for photosensitized reactions in natural waters (14,33-35). The concentration of dissolved organic carbon, which is predominantly humic substances (36))in the Skidaway River was 4.8 mg L-l (Table I). The principal products of photolysis of 2,4dichlorophenol in .distilled water were three isomers of chlorocyclopentadienic acid (11). We have tentatively identified similar photoproducts in 2,4-dichlorophenol photolysis in estuarine water (R. F. Lee and H.-M. Hwang, unpublished data). Pentachlorophenol had a lower photolysis rate in estuarne water than in distilled water. This is in agreement with the work of Miille and Crosby who showed that the decreased photolysis rate of pentachlorophenol in seawater relative to distilled water was due to photonucleophilic interaction of pentachlorophenol with chloride ion (25). We found out that chloride ihhibited pentachlorophenol photolysis but not the photolysis of dichlorophenol or trichlorophenol (data not shown). The number of chlorines and their position on the ring affect the interaction of chloride ion with the different chlorophenols (D. G. Crosby, personal communication). In summary, photolysis was found to be the primary transformation process in surface estuarine water for polychlorinated phenols. For phenol and monochlorophenol, microbial degradation was the primary transformation process. The winter decrease in surface irradiance and lower water temperatures resulted in a decrease in both photolysis and microbial degradation rates.
Acknowledgments We are greatly indebted to R. G, Zepp of the U.S.EPA in Athens for his advice and assistance in this work. We thank S. Keeran for his technical assistance and helpful discussions. We thank Dannah McCauley and Ching-Lee Hwang for their excellent typing work. Registry No. Phenol, 108-95-2; p-chlorophenol, 106-48-9; 2,4-dichlorophenol, 120-83-2; 2,4,5-trichlorophenol, 95-95-4; pentachlorophenol, 87-86-5.
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Received for review November 11,1985. Accepted April 18,1986. This work is a result of research sponsored by N O A A Office of Sea Grant, U. S. Department of Commerce, under Grant N A 8 4 A A -D -00072.
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