Apr 13, 1990 - Genthner, B. R. S., W. A. Price II, and P. H. Pritchard. 1989. ... 1987. Anaerobic biodegradation of monochlorophenols. Envi- ron. Technol. Lett.
Vol. 56, No. 11
APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Nov. 1990, p. 3255-3260
0099-2240/90/113255-06$02.00/0 Copyright ) 1990, American Society for Microbiology
Chlorophenol Degradation Coupled to Sulfate Reduction MAX M. HAGGBLOM' AND L. Y.
Departments of Microbiology' and Environmental Medicine,2 New York University Medical Center, 550 First Avenue, New York, New York 10016 Received 13 April 1990/Accepted 10 August 1990
We studied chlorophenol degradation under sulfate-reducing conditions with an estuarine sediment inoculum. These cultures degraded 0.1 mM 2-, 3-, and 4-chlorophenol and 2,4-dichlorophenol within 120 to 220 days, but after refeeding with chlorophenols degradation took place in 40 days or less. Further refeeding greatly enhanced the rate of degradation. Sulfate consumption by the cultures corresponded to the stoichiometric values expected for complete oxidation of the chlorophenol to CO2. Formation of sulfide from sulfate was confirmed with a radiotracer technique. No methane was formed, verifying that sulfate reduction was the electron sink. Addition of molybdate, a specific inhibitor of sulfate reduction, inhibited chlorophenol degradation completely. These results indicate that the chlorophenols were mineralized under sulfidogenic conditions and that substrate oxidation was coupled to sulfate reduction. In acclimated cultures the three monochlorophenol isomers and 2,4-dichlorophenol were degraded at rates of 8 to 37 ,umol liter-' day-'. The relative rates of degradation were 4-chlorophenol > 3-chlorophenol > 2-chlorophenol, 2,4-dichlorophenol. Sulfidogenic cultures initiated with biomass from an anaerobic bioreactor used in treatment of pulp-bleaching effluents dechlorinated 2,4-dichlorophenol to 4-chlorophenol, which persisted, whereas 2,6-dichlorophenol was sequentially dechlorinated first to 2-chlorophenol and then to phenol.
Contamination of the environment by chlorinated aromatic compounds has been the subject of increased concern in the last few years. Chlorinated phenols are common environmental contaminants; they have been extensively used as biocides, mainly as wood preservatives (26). Chlorinated phenols and other chlorinated phenolic compounds are also formed as by-products when chlorine is used for bleaching of pulp (22) and for disinfection of drinking water and wastewater containing phenols (1, 7). They are also formed during combustion of organic matter (2) and as biological breakdown products of chlorophenoxyacetic acid herbicides (12, 27). A range of chlorinated organic compounds including chlorophenols can be produced by biologic chlorination as well (24). In water, chlorophenols sorb onto particulate material and, if not degraded, eventually end up in sediments. Chlorinated phenolics have been found to accumulate in freshwater and marine environments where they may attain concentrations of tens of milligrams per kilogram of dry sediment (28, 38). In anoxic sediments, nitrate, sulfate, or carbonate may serve as a terminal electron acceptor for degradation of organic material. Anaerobic degradation of chlorophenols has mainly been studied under methanogenic conditions (5, 6, 10, 11, 15, 16, 20, 23, 33). These studies with freshwater sediments, soil, and sewage sludge as inoculum have shown that degradation of chlorophenols is initiated by reductive dechlorination, with complete mineralization to CO2 and CH4 observed in some cases. In marine environments, sulfate reduction is the major electron sink during anaerobic degradation of organic matter. In a marine sediment, it accounted for >50% of the mineralization of organic matter (35), while in a salt marsh environment the sulfate-mediated oxidation of organic matter was 12 times that of 02-mediated oxidation (14). The marine environment is a rich source of biologically produced *
halogenated compounds (24), suggesting that bacteria capable of (aerobic or anaerobic) dehalogenation could evolve in such habitats. Recently, anaerobic degradation of a naturally occurring halophenol, 2,4-dibromophenol, was observed in marine sediments (19). A number of reports indicate that sulfate appears to inhibit anaerobic degradation of chlorophenols (11, 33), but there is also evidence of chlorophenol degradation in the presence of sulfate (9) or during sulfate reduction (21). Whether degradation of chlorophenols can be linked to sulfate reduction has yet to be established. In this paper, we demonstrate that the degradation of chlorophenols can be coupled to sulfate reduction, as observed both with sediment from a polluted intertidal strait and with inoculum from a bioreactor shown previously to dechlorinate chlorolignin. MATERIALS AND METHODS Establishment of cultures. Strict anaerobic techniques were followed throughout the study. A sediment sample from an estuarine intertidal strait (East River, New York City) was used as inoculum. Sediment slurries were added as a 10% (vol/vol) inoculum, and a freshwater or saline sulfate medium was added to a total volume of 50 ml to deoxygenated 65-ml serum bottles. The freshwater medium was modified from Widdel (Ph.D. thesis, University of Gottingen, Gottingen, Federal Republic of Germany, 1980) and contained the following (in grams per liter): NaCl, 1.17; MgC12 6H20, 0.41; KCI, 0.3; CaCl2, 0.11; NH4Cl, 0.27; KH2PO4, 0.2; Na2SO4, 2.84; NaHCO3, 2.52; NaMoO4, 0.018 mg/liter; Na2S, 1.5 mM. The medium was supplemented with trace elements (37) and vitamins (Widdel, Ph.D. thesis), pH 7.2, with resazurin as a redox indicator. The saline sulfidogenic medium contained 23.0 g of NaCl and 1.0 g of MgCl2 per liter (otherwise as above), based on the measured salinity of the East River. The headspace gas was N2 (70%)-CO2 (30%). All bottles were sealed with butyl rubber stoppers and aluminum crimp seals. 2-Chlorophenol (2-CP), 3-chlorophenol (3-CP), 4-chlorophenol (4-CP), or 2,4-dichlo-
Corresponding author. 3255
rophenol (24-DCP) (Aldrich Chemical Co., Milwaukee, Wis.) was added to an initial concentration of 0.1 mM. The cultures were established in duplicate with background (no substrate added) and sterile (autoclaved three times on consecutive days) controls. The cultures were incubated statically at 30°C, in the dark. Another set of experiments was set up with inoculum from a laboratory-scale anaerobic fluidized-bed bioreactor used for treatment of pulp bleaching effluents (M. Haggblom and M. Salkinoja-Salonen, Water Sci. Technol., in press). Biomass from the bioreactor fluid was collected by centrifugation, washed, and suspended to 1/10 of the original volume in a phosphate buffer; 2 ml was added as inoculum to 50 ml of sulfate medium (described above). Establishment of cultures was otherwise as described above. The cultures were fed 1.0 mM propionate as an auxiliary substrate and incubated for 2 weeks prior to feeding with chlorophenols. 2,6-Dichlorophenol (26-DCP) (Aldrich Chemical Co.) and 24-DCP were fed to an initial concentration of 0.1 mM. The cultures were set up in duplicate with background and sterile controls and incubated as above. Analysis. Gas and liquid samples were taken for periodic analysis with sterile syringes, which had been deoxygenated with N2-C02. CH4 in the headspace gas was analyzed as described previously (4), with a gas partitioner (model 1200; Fisher Scientific Co., Springfield, N.J.) equipped with a thermal conductivity detector. Sulfate was analyzed by an indirect titration as follows (13). Sulfide was first removed by precipitation as ZnS. Sulfate was then precipitated as barium sulfate in acid EDTA solution, filtered, and dissolved in an excess of EDTA at high pH, and the excess EDTA was titrated with MgCl2. Chlorophenols were quantified by high-performance liquid chromatography. Prior to analysis, the samples (0.3 to 0.5 ml) were acidified with 10 pAl of 1 N HCl, centrifuged, and filtered (0.45 tLm). Analysis was performed with a Beckman 332 LC chromatograph (Beckman Instruments, Palo Alto, Calif.) equipped with a Spherisorb C-18 column (250 by 4.6 mm; Supelco Inc., Bellefonte, Pa.), with UV detection at 280 nm, and using a solvent system of methanol (60%, vol/vol)water (38%, vol/vol)-acetic acid (2%, vol/vol) at a flow rate of 1 ml/min. Determination of sulfide formation. The reduction of sulfate to sulfide was determined by using a modified radiotracer technique (17, 18). A 4-CP-degrading culture obtained through repeated transfers into fresh medium and refeeding with chlorophenol was used. The culture was split into 5-ml subcultures in 10-ml vials, and 5 nCi of NaSO4 (43 Ci/mg, carrier-free, 99% radionuclidic purity; ICN Radiochemicals, Irvine, Calif.) was added. Replicate cultures were fed chlorophenol twice to a total of 0.5 mM and incubated for 2 weeks. Lactate was used as a nonselective substrate for sulfate reducers as an active control. Molybdate-inhibited and unfed cultures served as controls. When the chlorophenol had been degraded, the cultures were acidified with 25% HCI, which releases sulfide as H2S gas. H2S was then driven off by flushing the vessel with argon for 30 min and collected in a series of two Zn acetate (2%, wt/vol) traps, where sulfide was precipitated as ZnS. An
E 0.06-C 0
IU* 100 150 time (days)