Environ. Sci. Technol. 1991, 25, 446-449
93; American Chemical Society: Washington, DC, 1979; DD 215-298. Benjamin, M. M. Ph.D. Dissertation, Stanford University, 1979. Vanek, K.; Jedinakova, V. J . Colloid Interface Sei. 1986, 1I I , 276-219. Kinniburgh, D. G.; Jackson, M. L. In Adsorption of Inorganics at Solid-Liquid Interfaces; Anderson, M. A., Rubin, A. J., Eds.; Ann Arbor Science: Ann Arbor, MI, 1981. Kinniburgh, D. G.; Baker, J. A.; Whitfield, M. J . Colloid Interface Sei. 1983, 95, 370-384. Snedecor, G. W.; Cochran, W. G. Statistical Methods, 6th ed.; The Iowa State University Press: Ames, IA, 1976. Westall, J . C. In Aquatic Surface Chemistry; Stumm, W., Ed.; John Wiley and Sons: New York, 1987; Chapter 1.
(48) Dzombak, D. A.; Morel, F. M. M. J. Colloid. Interface Sei. 1986, 112, 588. (49) Dzombak, D. A.; Morel, F. M. M. J . Hydraul. Eng. 1987, 113, 430. (50) Goldberg, S.; Traina, S. J. Soil Sei. SOC.A m . J . 1987, 51, 929-932. (51) Chu, S.-Y.; Sposito, G. Soil Sci. Soc. A m . J . 1981, 45, 1084-1089.
Received for review January 18, 1989. Revised manuscript received June 18,1990. Accepted October 10,1990. This research was funded by the Electric Power Research Institute, Inc. (EPRT) under Contract RP2485-03, Chemical Attenuation Studies, to Battelle, Pacific Northwest Laboratories.
Brominating Activity of the Seaweed Ascophy//um nodosum: Impact on the Biosphere Ron Wever," Michiel G. M. Tromp, Bea E. Krenn, Abdoeljallal Marjani, and Mauritz Van To1 E. C. Slater Institute for Biochemical Research and Biotechnological Center, University of Amsterdam, Plantage Muidergracht 12, 1018 TV Amsterdam, the Netherlands
Macroalgae are an important source of volatile halogenated organic compounds, such as bromoform and dibromomethane. The mechanism by which these compounds are formed is still elusive. We report that the brown seaweeds Laminaria saccharina, Laminaria digitata, Fucus uesiculosis, Pelvetia canaliculata, and Ascophyllum nodosum and the red seaweeds Chondrus crispus and Plocamium hamatum contain bromoperoxidases. The intact plants are able to brominate exogeneous organic compounds when HzO, and Br- are added to seawater. Further, we show that the brominating activity of the brown macroalga A . nodosum, which contains a vanadium bromoperoxidase located on the thallus surface, occurs when the plant is exposed to light and not in the dark. The rate of bromination of exogenous organic compounds in seawater by this plant is 68 nmol (g of wet alga1-l h-l. HOBr is a strong biocidal agent and we propose that the formation of HOBr by this seaweed is part of a host defense system.
Introduction There is great current interest in the formation of organohalides produced by both anthropogenic activities and biological sources. Recently, it has been suggested (1) that chemical reactions occur in the troposphere at polar sunrise in the Arctic. In one of these processes ozone destruction occurs a t ground level, which may be connected to the formation of bromoform and aerosol enrichment with bromine of biological origin (2). According to Sturges and Barrie (2), inorganic bromine in Arctic aerosols may originate from photochemical conversion of marine organic bromine. Measurements ( 3 ) of atmospheric bromoform a t Point Barrow, AK, show that the presence of this compound is seasonal: bromoform concentrations are maximal in winter and minimal in summer. Similarly, the bromine content of Arctic aerosols at Point Barrow, Alert (Canadian Arctic), and Spitsbergen shows an annual sharp maximum between February and May at concentrations that are the highest in the world ( 2 , 4 , 5 ) . There is substantial evidence that the source of the bromine-containing particles and bromoform is biogenic and it has been speculated that the formation is due to the enzymic activity of vanadium 446
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bromoperoxidases present in brown seaweeds that grow in the Arctic Ocean (6). Indeed, Dyrssen and Fogelqvist (7, 8) showed that bromoform is present in the Arctic Ocean near Spitsbergen and the data clearly indicate that algal belts are responsible for its production. It is wellknown that brown seaweeds (Ascophyllum nodosum, Fucus uesiculosis), red seaweeds (Gigartina stellata), and marine green seaweeds (Enteromorpha linza, Ulva lacta) release large quantities of brominated methanes such as bromoform and dibromomethane into the marine ecosystem (9). Recently, a group of vanadium-containing bromoperoxidases has been discovered in a number of brown seaweeds (Fucus distichus, Alaria esculenta, Laminaria saccharina, Chorda filum) and in red seaweeds (Ceramium rubrum and Corallina pilulifera) (10-14). In the presence of hydrogen peroxide and bromide these enzymes (15) produce free hypobromous acid and bromine. The peroxidases are assumed to be involved in the biosynthesis of the halogenated metabolites by bromination of nucleophilic acceptors. As has been shown by Theiler et al. (16) for the marine red alga Bonnemaisonia hamifera and by Burreson et al. (17) for the red alga Asparagopsis taxiformis, the acceptors are probably carboxy methyl ketones, which are brominated by purified bromoperoxidase and then decay to form brominated methanes. However, whether similar acceptors also exist in brown or other red seaweeds is not known and other pathways may exist. The brown seaweed A . nodosum (knotted wrack), which grows in large quantities in the North Atlantic and western Russian polar seas (18), contains two vanadium bromoperoxidases (19). One is located inside the so-called fruiting bodies and its presence shows a seasonal cycle (6). The other enzyme is located on the surface of the fruiting bodies and may be extracellular (19). This raises the interesting possibility that these seaweeds also release free hypobromous acid and bromine in seawater. Reaction of these bromine species with dissolved organic matter in seawater may well result in the formation of bromoform and other brominated compounds, and thus the origin of bromoform may be an indirect result of the formation of H O B r by seaweeds. In fact, Sauvageau (20) already re-
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1
0.0 0
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500 X(nm1
Figure 1. Bromination of phenol red by A . nodosum in natural seawater. About 50 g of the seaweed was incubated in 200 mL of seawater (25 "C)to which 25 p M phenol red was added and the seaweed was either exposed to sunlight or kept in the dark. A, start of the incubation; B, 0.5 h; C, 1 h; D,2 h; E, 3 h exposed to sunlight.
ported in 1926 that the red seaweeds Antithamnionella sarniensis and Antithamnion Plumula were able to release HOBr and Br, in seawater. To detect the presence of bromine he used fluorescein, which will be converted to eosin, a tetrabrominated compound. However, his results were published in a French journal and have apparently been overlooked.
Experimental Section Phenol red was used as an indicator of the formation of HOBr, since bromination of this dye leads to bromophenol blue, which has a typical absorption band a t 592 nm at neutral pH. Bromination of this dye by isolated bromoperoxidase was studied by us (21). Details of haloperoxidase-catalyzed bromination of phenol red can also be found in ref 22. Fresh A. nodosum and seawater were collected in the period August to November 1989 or in September 1990 near Kornwerderzand; L. saccharina, F. uesiculosis, and Chondrus crispus were collected in March 1990 along the dike of the Oosterschelde, the Netherlands; Peluetia canaliculata and Laminaria digitata were collected in April 1990 hear Oban, Scotland; Plocamium hamatum was collected in June 1989 on Heron Island, The Great Barrier Reef, Australia. The algae were transported in seawater to the laboratory and the seawater was centrifuged at 2500g to remove particles. The absorption coefficients used in calculating the amount of (tetra)bromophenol blue and phenol red were 67.4 mM-' cm-l at 592 nm and 19.7 mM-l cm-l at 433 nm (211, respectively. A t the laboratory, 20, 30, or 50 g of the alga A. nodosum (in total, six plants) was placed in beakers filled with 200 mL of seawater (25 OC) containing 25 pM phenol red and these glass beakers were either kept in the dark or exposed to direct sunlight. Aliquots (2.25 mL) were taken and 0.25 mL of morpholinoethanesulfonic acid (pH 6.5) was added to a final concentration of 0.1 M to bring the samples to the same pH. To test the localization of bromoperoxidase, -30 g of A. nodosum was incubated in the dark in 110 mL of seawater (25 O C ) containing 20 pM phenol red to which 2 mM H 2 0 2and 100 mM KBr were added. The other seaweeds (10-40 g) were incubated in 200 mL of the same medium. The rate of formation of bromophenol blue was calculated from the absorbance increase at 592 nm after incubation for 1 h. Results and Discussion When fresh A. nodosum was incubated with 25 pM phenol red in natural seawater, bromination of phenol red (Figure 1) was clearly observed when the sample was allowed to stand in full sunlight. In the dark no reaction occurred (not shown), and apparently this phenomenon
20
40
timin)
60
Flgure 2. Bromination of phenol red by A . nodosum in the presence of exogenous H,O, and bromide. About 30 g of the seaweed was incubated in 110 mL of seawater (25 "C)to which 20 pM phenol red was added. The experiments were carried out in the dark. 0, no additions; 0 , 2 mM H,O,; 0,2 mM H20, and 100 mM KBr.
is linked to, or triggered by, a photochemical process. However, the alga is clearly involved in this, since upon illumination by sunlight of seawater only, no bromination of phenol red was detectable (not shown). From the changes in absorbance at 433 and 592 nm it is possible to calculate the amount of phenol red that disappeared and the amount of bromophenol blue that was formed. Comparison of the spectra obtained at the start of the illumination (trace A) and that after 3 h (trace E) shows a good correlation between the amount of phenol red (9.9 pM) converted into bromophenol blue (9.0 pM). The amount of phenol red brominated was determined for six different plants by measuring the difference in absorbance a t 592 nm between control samples, which were kept in the dark, and samples that were exposed to sunlight for 1 h. This yielded a value of 16.9 f 8.7 nmol of bromophenol blue formed (g of wet weight of alga)-l h-l. As shown by the De Boer and Wever (15), the vanadium bromoperoxidases produce free hypobromous acid and bromine. Therefore, it is highly likely that bromination of the dye by the seaweed is due to reaction with HOBr released by the intact plants. The rate of formation of bromophenol blue found would therefore correspond to a rate of HOBr formation of 68 nmol g-' h-l. As shown (19),the vanadium bromoperoxidase located on the thallus surface can be removed by rinsing the alga for a short period in 0.2 M Tris-HC1 (pH 8.3). It is likely that alga treated in this manner will not be able to form HOBr. Indeed, bromination of phenol red was not observed. We also tested whether light from a Xenon lamp (150 W) was able to induce HOBr formation. Bromination of phenol red occurred, but the rate (7 nmol g-' h-l) was lower than that obtained in full sunlight, apparently due to lower light intensity. We further tested whether the bromoperoxidase on the thallus surface of A. nodosum was accessible to substrates added externally. Figure 2 illustrates the effect of added H z 0 2and bromide on the rate of bromination in natural seawater catalyzed by the seaweed in the dark. When no HzOz was added to the medium, only a small amount of bromophenol blue was found. However, when 2 mM H202 was present, bromination of phenol red occurred, the rate of which greatly accelerated when 100 mM bromide was added to the seawater. It should be noted that seawater contains 1mM bromide (23);the K , for bromide of the bromoperoxidase at the thallus surface is 0.4 mM at pH 4.7, increasing to 6.7 mM at pH 7.2 (19). Thus, the enzyme on the thallus surface operates below V,,, conditions, which would explain the increase in rate when 100 mM bromide was added (Figure 2).
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Table I. Rate of Bromination of Phenol Red by Macroalgae"
species L. saccharina L. digitata A . nodosum P. canaliculata F. uesiculosis C. crispus P. hamatum
nmol of bromophenol blue g-' h-' 41 123
63 56 6 26
+*
"The rate was measured after addition of 2 mM H,O, and 100 mM Br-. *The actual rate was not determined.
To test whether other seaweeds (L. saccharina, L. digitata, F. uesiculosis, P. canaliculata, Plocamium hamatum, and C. crispus) also contain bromoperoxidases that are accessible to added substrates and are able to brominate phenol red, 2 mM H 2 0 2and 100 mM Br- were added to these seaweeds in seawater. Table I summarizes the results of this experiment. The rate of bromination catalyzed by the intact plants is of the same order of magnitude; only the F. vesiculosis shows a lower rate. Thus, it is likely that the bromoperoxidases are also present extracellularly on the thallus surfaces of these plants. However, unlike the observation for A . nodosum, formation of Br, and HOBr by these seaweeds was not induced by sunlight. In conclusion, we show here for the first time that the bromoperoxidase in A . nodosum is able to brominate exogeneous organic compound in seawater and that its brominating activity can be triggered by light or addition of substrates. Gschwend et al. (9) reported that the brown seaweed A . nodosum released -4500 ng of CHBr, (g of dry alga)-l day-'. The biosynthetic pathway by which CHBr, is formed by this alga is not known a t present. For the red seaweed Bonnemaisonia hamifera it has been shown (16) that halogenated metabolites are being produced via bromination, mediated by a bromoperoxidase, of ketones yielding halogenated ketones. These ketones are unstable and decay to form volatile brominated methanes (17). It may be that such a pathway to produce CHBr, is also operative in A . nodosum. Since the bromoperoxidase is located on the thallus surface, this requires, however, that ketones are present a t or near the surface. In line with this, Gschwend et al. (9) concluded that the brominated compounds are synthesized near the surface of A . nodosum. A more likely possibility is, we feel, that HOBr is just released in seawater and reacts with dissolved organic matter (24) to form unstable brominated compounds, which decay to give rise to bromoform. This is supported by the observation (25) that chlorination of dissolved organic material in freshwater leads to the formation of chloroform and other volatile metabolites. Further, Helz and Hsu (26) demonstrated direct formation of CHBr, upon chlorination of contaminated coastal water. Chlorination of seawater will rapidly oxidize bromide in seawater to HOBr, which reacts extremely rapidly with organic matter (24). This pathway of bromoform production requires, however, that the rate of HOBr formation observed in our study would be higher than the reported CHBr, formation by these algae since not all HOBr will end up in bromoform. The value reported (9) corresponds to 17.8 nmol of CHBr, (g of dry weight)-' day-'. Assuming that the natural photoperiod used in the study was 10 h and that the dry weight of this alga is -50% of its wet weight, this rate would correspond to -0.9 nmol of CHBr, (g of wet weight)-' h-l. Since a t least 3 nmol of HOBr is required to form 1 nmol of bromoform, this would corre448
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spond to 2.7 nmol of HOBr (g of wet weight)-'. This value is -4.0% of the value found for the alga exposed to full sunlight in this study. From the rate of release of halometabolites by several algae and a global biomass (27) of lo1, g, Gschwend et al. (9) estimated an annual global input in the biosphere of lo4 tonslyear and concluded that the macroalgae may be an important source of bromine-containing material released in the atmosphere. As we demonstrated here, formation of HOBr by the vanadium bromoperoxidases present in these seaweeds may be a major step in the formation of these volatile halogenated compounds. Further, the amounts of HOBr produced yearly are substantial. Assuming a biomass of 2 X los g of A . nodosum along the Afsluitdijk (length 30 km), an input of -1.8 X lo6 g of HOBr in the Waddenzee can be estimated due to the activity of this seaweed. It is conceivable that some fraction of HOBr and Brz escapes the water and is directly released to the air, resulting in bromine-containing particles. It may well be that the annual sharp maximum observed in bromine content of Arctic aerosol, which peaks every year just after Arctic dawn ( 2 , 5 ) ,is related to photosynthetic activity of seaweeds present in the Arctic Ocean. As we have shown, the brown seaweed A . nodosum releases HOBr when exposed to light. Not all seaweeds show this phenomenon. However, in the species investigated, bromoperoxidases are found on the thallus surface which are accessible to H,Oz and Br-. Surface waters may contain up to 1 pM HzOz (28), and it is conceivable that the bromoperoxidases in these plants are using HzO2that is generated photochemically or biologically (29). The function of formation of HOBr by seaweed is elusive. However, HOBr, like HOC1, is a strong biocidal agent and we propose that the generation of the agent by the seaweed is part of a host defense system by which the alga prevents bacterial and fungal infections or prevents herbivore feeding. Such a system is not without precedent in eukaryotes. Haloperoxidase-mediated antimicrobial systems have been shown to be operative in neutrophils (30) and eosinophils (31). These white blood cells contain myeloperoxidase and eosinophil peroxidase, respectively, which are known to catalyze the peroxidation of chloride and bromide into hypochlorous acid and hypobromous acid. Registry No. HOBr, 13517-11-8;bromoperoxidase, 69279-19-2.
Literature Cited (1) Barrie, L. A.; Bottenheim, J. W.; Schnell, R. C.; Crutzen, P. J.; Rasmussen, R. A. Nature 1988, 334, 138. (2) Sturges, W. T.; Barrie, L. A. Atmos. Enuiron. 1988,22,1179. (3) Cicerone, R. J.; Heidt, L. E.; Pollock, W. H. J . Geophys. Res. 1988, 93, 3745. (4) Berg, W. W.; Heidt, L.E.; Pollock, W.; Sperry, P. D.; Cicerone, R. J. Geophys. Res. L e t t . 1984, 11, 429. (5) Berg, W. b'.;Sperry, P. D.; Rahn, K. A,; Gladney, E. S. J. GeoDhvs. Res. 1983, 88, 6719. (6) We;er; R. Nature 1988; 335, 501. (7) Dyrssen, D.; Fogelqvist, E. Oceanol. Acta 1981, 4 , 313. (8) Fogelqvist, E. J . Geophys. Res. C 1985, 90, 9181. (9) Gschwend, P. M.; MacFarlane, J. K.; Newman, K. A. Science 1985, 227, 103. (10) De Boer, E.; Van Kooyk, Y.; Tromp, M. G. M.; Plat, H.; Wever, R. Biochim.Biophys. Acta 1986,869, 48. (11) De Boer, E.; Tromp, M. G. M.; Plat, H.; Krenn, B. E.; Wever, R. Biochim. Biophys. Acta 1986,872, 104. (12) Krenn, B. E.; Plat, H.; Wever, R. Biochim. Biophys. Acta 1987, 912, 287. (13) Krenn, B. E.; Izumi, Y.; Yamada, H.; Wever, R. Biochim. Biophys. Acta 1989, 998, 63. (14) Wever, R.; Olafsson, G.; Krenn, B. E.; Tromp, M. G. M. Abstracts, 32nd IUPAC Congress; Stockholm, 1989; p 31.
Environ. Sci. Technol. 1991, 25, 449-455
(15) De Boer, E,; Wever, R. J . Biol. Chem. 1988, 263, 12326. (16) Theiler, R.; Cook, J. C.; Hager, L. P.; Siuda, J. F. Science 1978,202, 1094. (17) Burreson, B. J.; Moore, R. E.; Roller, P. P. J . Agric. Food Chem. 1976, 24, 856. (18) Zenkevitch, L. Biology of the Seas of the U S S R ; Interscience Publishers: New York, 1963; p 955. (19) Krenn, B. E.; Tromp, M. G. M.; Wever, R. J . Biol. Chem. 1989, 264, 19287. (20) Sauvageau, C. Bull. Stat. Biol. Arc. 1926, 23, 5. (21) De Boer, E.; Plat, H.; Tromp, M. G. M.; Franssen, M. C. R.; Van Der Plas, H. C.; Meijer, E. M.; Schoenmaker, M. E.; Wever, R. Biotechnol. Bioeng. 1989, 30, 607. (22) Neidleman, S.L.; Geigert, J. In Biohalogenation: Principles, Basic Roles and Applications; Ellis Horwood Ltd: Chichester, 1986; p 70. (23) Riley, J. P.; Chester, R. In Introduction to Marine Chemistry; Academic Press: London and New York, 1971; p 60. (24) Jaworske, D. A.; Helz, G. R. Environ. Sci. Technol. 1985, 19, 1188. (25) Scully, F. E.; Howell, G. D.; Kravitz, R.; Jewell, J. T.;Hahn, V.; Speed, M. Environ. Sci. Technol. 1988, 22, 537.
Helz, G. R.; Hsu, R. Y. Limnol. Oceanogr. 1978,23, 859. Waaland, R. J. In T h e Biology of Seaweeds; Lobban, C. S., Wynne, M. J., Eds.; University of California Press: Berkeley and Los Angeles, 1981; pp 726. Petasne, R. G.; Zika, R. G. Nature 1987, 325, 516. Palenik, B.; Zafiriou, 0. C.; Morel, F. M. M. Limnol. Oceanogr. 1981, 32, 1365. (30) Foote, C. S.; Goyne, T. E.; Lehrer, R. I. Nature 1983,301,
715. (31) Weiss, S. J.; Test, S. T.; Eckmann, C. M.; Roos, D.; Regiani, S. Science 1986, 234, 200.
Received for review M a y 3, 1990. Revised manuscript received September 24,1990. Accepted October 1, 2990. W e thank the Dutch State Mines, Geleen, for their support; we greatly acknowledge the gift by Hewlett-Packard, the Netherlands, of a 8452A spectrophotometer and the support of Prof. Dr. C. J . Hawkins (University of Queensland, Queensland, Australia). W e thank Dr. D. Parry for his help in collecting the alga P. hamatum. These investigations are partly supported by the Netherlands Foundation for Chemical Research (S.O.N.).
Precipitation of Jarosite Compounds as a Method for Removing Impurities from Acidic Wastes from Chemical Coal Cleaning Glenn A. Norton,* Rlchard G. Richardson, and Richard Markuszewski Ames Laboratory, I o w a State University, Ames, I o w a 5001 1
Audrey D. Levine Department of Civil and Construction Engineering, I o w a State University, Ames, I o w a 5001 1
Precipitation of jarosite compounds to remove Na, Fe, and from spent acid solutions from a chemical coal-cleaning process was studied in relation to reaction time and pH. Although Fe and S042-could be removed effectively from model solutions a t p H values of 1.5-2.3, optimum Na removal was possible only within the narrow p H range of 1.4-1.6. Maximum precipitate yields were obtained within -6 h a t both 80 and 95 "C, with a t least 80% of the Fe and S042-and -60-7570 of the Na removed. An additional benefit of precipitation of jarosite compounds to remove impurities from spent acid streams is that these compounds have low solubilities in water and are therefore attractive from a waste disposal perspective. Based on this study, the feasibility of cleaning spent acid from a chemical coal-cleaning process by precipitating jarosite compounds has been demonstrated. Introduction
Conventional coal-cleaning technologies can remove much of the inorganic sulfur in coals by using physical methods based on differences in density or surface properties between minerals and coal macerals. However, organic sulfur comprises typically 40-6070 of the total sulfur in bituminous coals, and it cannot be removed effectively by physical separations. Thus, chemical treatments are necessary to remove the organic sulfur from coal. Although physical cleaning methods have been used for many years, chemical coal cleaning is still in the developmental stages. The TRW Gravimelt process, also known as the molten caustic leaching (MCL) process, is one chemical cleaning procedure that has been studied in detail in recent years (1-6). A general schematic diagram of the MCL process is shown in Figure 1. First the coal is leached with molten caustic (NaOH or a NaOH/KOH mixture) at 300-400 "C 0013-936X/91/0925-0449$02.50/0
for 1-3 h. The reacted coal/caustic cake is then washed consecutively with water, a dilute sulfuric acid solution, and again with water. This process produces a low-sulfur and low-ash coal product. The spent acid from the washing stage of the MCL process contains substantial amounts of Na, Fe, and SOf impurities and may require treatment to remove these impurities before it can be recycled back into the acidwashing step or discharged to the environment. One option for treating the spent acid stream is to precipitate the impurities as double salts in the jarosite family of compounds. The theoretical composition for jarosite compounds is MFe3(S04)2(0H)6, where M is a monovalent cation from the group H30+,Na+, K+, Rb+, Ag+, NH4+, T1+, 1/zPb2+,or 1/2Hg2+(7,8). Alkali jarosite compounds are crystalline, can be easily dewatered and dried, and may be resistant to leaching, thus providing a stable compound that can be disposed of without additional treatment (9). Little information is available on the solubility of jarosite compounds. However, according to one literature source, the solubility of natrojarosite in water is -0.02 g/L (10). In this study, the feasibility of removing impurities from acid solutions by precipitation of jarosite (KFe3(S04)2(OH),) and natrojarosite (NaFe3(S04),(OH),)was evaluated. The goal of this work was to determine general operating guidelines for precipitating jarosite compounds from spent acid solutions typical of those produced in the MCL process. The effects of varying the reaction time, temperature, and pH on the removal of Na, Fe, and SO-: were examined. The precipitate yield, chemical and mineralogical composition of the precipitate, and dissolved ions present in the solution at the completion of each test were studied with model solutions. The results obtained from studies with model solutions were applied to spent acids produced during laboratory
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