Environ. Sci. Technol. 2000, 34, 3384-3388
Mineralization of RDX by the White Rot Fungus Phanerochaete chrysosporium to Carbon Dioxide and Nitrous Oxide TAMARA W. SHEREMATA AND JALAL HAWARI* Biotechnology Research Institute, National Research Council, 6100 Royalmount Avenue, Montreal, Quebec, H4P 2R2 Canada
Biodegradation of hexahydro-1,3,5-trinitro-1,3,5-triazine (RDX) in liquid cultures (initially at 62 mg/L) was studied using the white rot fungus Phanerochaete chrysosporium. With RDX as the main source of nitrogen, complete disappearance occurred after 60 d. The major products of RDX transformation were CO2 and N2O, and both gases appeared after a 2-d lag period. Following 60 d, an average of 52.9% [UL-14C]-RDX was mineralized to CO2, 10.7% was taken up as biomass by the fungi, and 28.3% remained in the aqueous phase as unidentified metabolites. Of the nitrogen in RDX, 62.0% was transformed to N2O. Transformation of ring-labeled [15N]-RDX, with subsequent analysis by GC-MS, indicated that the N2O was composed of one nitrogen atom from the RDX ring and the other nitrogen from one of the nitro group substituents. Oxidation of RDX was correlated with manganese peroxidase (MnP) enzyme activity (lignin peroxidase (LiP) activity was absent). Traces of 1-mononitroso-3,5-dinitro-1,3,5-triazine were evident throughout the course of the experiment. Results of this study provide new information regarding N2O as a major product of RDX mineralization. Quantification of N2O at sites contaminated with RDX may be an important parameter for monitored natural attenuation.
Introduction Hexahydro-1,3,5-trinitro-1,3,5-triazine or Royal Demolition Explosive (RDX, Figure 1) is an energetic compound that is commonly used as a military explosive. Various commercial and military activities that include manufacturing, waste discharge, testing and training, demilitarization, and open burning/open detonation (OB/OD) have resulted in extensive RDX contamination of soil and groundwater (1). The toxicity of RDX to humans and mammals is well established (2). Hence, remediation of contaminated soil and groundwater is necessary. Bioremediation is a preferred method since it is less costly and does not produce dangerous emissions that are associated with physicochemical techniques (i.e., incineration or adsorption by granular activated carbon followed by alkaline hydrolysis) (3-5). The metabolic pathways of polynitroaromatics, such as 2,4,6-trinitrotoluene, have been extensively characterized in liquid culture and natural soil systems (6, 7). Until now there has been little information concerning the nonaromatic cyclic * Corresponding author telephone: (514)496-6267; fax: (514)4966265; e-mail:
[email protected]. 3384 9 ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 34, NO. 16, 2000
FIGURE 1. Molecular structures of RDX and MNX molecules. nitroamines, such as RDX, with respect to ring cleavage products and metabolic pathways (8-10). Recently, several metabolites and end products were identified in the biodegradation of RDX with an anaerobic sludge (11). Particularly, hexahydro-1-nitroso-2,5-dinitro-1,3,5-triazine (MNX, Figure 1) and hexahydro-1,3-dinitroso-5-nitro-1,3,5-triazine (DNX) were formed by the stepwise reduction of -NO2 in RDX. Methylenedinitramine (O2NNHCH2NHNO2) and bis(hydroxymethyl)nitramine ((OHCH2)2NNO2) product formation were attributed to enzymatic hydrolytic ring cleavage of RDX. The four metabolites identified above disappeared to produce N-containing products (N2O, NH4+, and traces of N2) as well as C-containing products (HCHO, CH3OH, HCOOH, and CO2). During the course of these experiments, 50-60% mineralization of RDX to CO2 occurred. Knowledge of RDX degradation products, as just described for anaerobic degradation of RDX (11), is necessary in order to understand the microbiology, biochemistry, and molecular biology of the biodegradation process. However, such information regarding the aerobic biodegradation of RDX is not available. Using Phanerochaete chrysosporium, Fernando and Aust (12) reported that 67% of RDX (initially at 0.02 mg/ L) was mineralized to CO2, 20.2% was recovered as watersoluble metabolites, and 96% of the RDX disappeared after 30 d. In contrast, using the same fungus, Bayman et al. (13) observed no mineralization of RDX (initially at 100 mg/L) with 22% RDX disappearance after 3 d. The differences observed by Fernando and Aust (12) and Bayman et al. (13) are likely due to the differences in initial RDX concentration and incubation times. Regardless, neither study identified metabolites or ring cleavage products of RDX. Binks et al. (9) reported two new intermediate products from RDX with the aerobic isolate Stenotrophomonas maltophilia. Using GCMS (EI), trace quantities of the chloride salt of methyleneN-nitroamino-N-acetoxyammonium chloride and methyleneN-(hydroxymethyl)hydroxylamine-N-(hydroxymethyl)nitramine were identified. The first product was assumed to be an impurity. The objective of the present study was to characterize the biotransformation of RDX in liquid culture by the white rot fungus P. chrysosporium. This microorganism was selected because of its nonspecific enzymatic activity that allows it to degrade a range of persistent or toxic compounds (14). As such, this fungus has been extensively studied for bioremediation. By characterizing some of the products from RDX biodegradation, the present study is a contribution toward these efforts.
Experimental Section Chemicals. RDX (> 99% purity) was provided by Defense Research Establishment Valcartier (Valcartier, PQ, Canada). Uniformly labeled [UL-14C]-RDX was synthesized and recrystallized to achieve chemical and radioactive purity of 99% and 97%, respectively (15). The specific activity of the radioactive compound was 28.7 µCi/mmol. The ring-labeled 10.1021/es000998y CCC: $19.00
Published 2000 by the Am. Chem. Soc. Published on Web 07/06/2000
[15N]-RDX (>98% purity) was synthesized similarly (16). The MNX was synthesized according to the methods of Brockman et al. (17). All other chemicals used were reagent grade. Strain. The fungal strain used was P. chrysosporium BKMF-1767 (ATCC 24725). The strain was maintained on malt agar slants (20 g of agar, 20 g of malt extract, and 1 g of yeast extract) at 37 °C (18). Spores were collected by washing the slants with distilled water. The spore suspension was stored at 4 °C. Microcosms for RDX Degradation. The medium (18) used for microcosms contained KH2PO4 (2 g L-1), CaCl2‚2H2O (0.14 g L-1), MgSO4‚7H2O (0.7 g L-1), FeSO4‚7H2O (0.07 g L-1), ZnSO4‚7H2O (0.046 g L-1), MnSO4‚H2O (0.035 g L-1), CuSO4‚ 5H2O (0.007 g L-1), glycerol (10 g L-1), disodium tartrate (2.3 g L-1), yeast extract (1 g L-1), thiamine (0.0025 g L-1), veratryl alcohol (0.067 g L-1), and commercial soybean phospholipids (NAT 89, 0.5 g L-1, Natterman Phospholipid GmbH, Koln, Germany). For each microcosm, 19 mL of medium was combined in a serum bottle (100 mL) with RDX and 1 mL of concentrated spores. The final concentration of RDX and spores was 62 mg/L and 2 × 106 spores/mL, respectively. Chemical controls were prepared as above with RDX, but with 1 mL of distilled water instead of concentrated spores. Biological controls were also prepared without RDX. All experiments were conducted in triplicate, and manipulations were carried out in a laminar flow biological hood to maintain aseptic conditions. The microcosms were incubated statically in the dark at 37 °C and were aerated three times a week. Mineralization experiments were prepared as above with 0.023 µCi [UL-14C]-RDX and a total RDX concentration of 62 mg/L. Each serum bottle was fitted with a small test tube containing 1 mL of 0.5 M KOH to trap liberated carbon dioxide. Microcosms with [UL-14C]-RDX were routinely sampled (every 2-3 days) for determination of 14CO2 in the KOH trap using a Tri-Carb 4530 liquid scintillation counter (model 2100 TR; Packard Instrument Company, Meriden, CT). At the end of the mineralization experiments, the microcosms were sacrificed so that a radioactive carbon balance could be determined. The aqueous phase was decanted and analyzed by liquid scintillation counting. The residual aqueous phase and the mycelial mat was combined with Soluene-350 (Packard Instrument Co., Downers Grove, IL), a fast-acting proteinaceous tissue solubilizer with a high water-holding capacity. The microcosms were left stationary for 3 d to allow for complete solubilization of the solid biomass. Samples of solubilized biomass were subsequently combined with Hionic-Fluor (Packard Instrument Co., Downers Grove, IL), a scintillation cocktail designed for large holding capacity of samples of high ionic strength. The homogeneous samples were analyzed by liquid scintillation counting, and the radioactivity taken up by the mycelial mat was obtained by difference (i.e., total radioactivity minus radioactivity in the residual aqueous volume). Enzyme Assays. Lignin peroxidase (LiP) activity was determined by monitoring the conversion of veratryl alcohol to veratryl aldehyde by hydrogen peroxide at 310 nm (19). Manganese peroxidase (MnP) activity was determined by monitoring the disappearance of vanillylacetone at 334 nm (20). Analytical Methods. RDX concentrations were determined by reversed-phase high-pressure liquid chromatography (HPLC) with a photodiode array (PDA) detector. The Waters (Waters Associates, Milford, MA) HPLC system consisted of a model 600 pump, a 717 plus autosampler, and a 996 PDA detector (λ ) 254 nm). A Supelcosil LC-CN column (25 cm × 4.6 mm, particle size 5 µm) was coupled with a temperature control module held at 35 °C. The methanol/ water gradient was at a flow rate of 1.5 mL/min. The initial solvent composition was 30% methanol and 70% water that was held for 8 min. Following this, a linear gradient was run
from 30 to 65% methanol over 12 min. The solvent ratio was changed to the initial conditions over 5 min and held for another 5 min. The system was outfitted with Millenium data acquisition software. The same HPLC detector system described above was used to analyze formic acid, a possible metabolite of RDX. In this case, an ion organic acid column (300 nm × 7.8 mm i.d.) was used (Phenominex, Torrance, CA). The column temperature was 30 °C, and the mobile phase was acidified water (pH 2.5) that consisted of concentrated sulfuric acid (95-98%) and demineralized water (degassed with helium). The flow rate was 0.4 mL/min, the injection volume was 100 µL, and UV detection was at 210 nm. An SRI 8610 GC (INSUS Systems Inc.) connected to a Supelco Porapack Q column (2m) was coupled with an electron capture detector (ECD) (330 °C) for N2O detection. Gas samples from the headspace of the microcosms were sampled using a gas-tight syringe for subsequent injection to the GC using helium as a carrier gas (21 mL/min) at 60 °C. Identification was confirmed by comparison with a reference compound. Following gas sampling, the headspace of the microcosms was replaced with air, and the N2O evolved was reported as a cumulative quantity. This is the same method used by Roy and Knowles (21) for N2O evolution during nitrification. The presence of N2O as a product of RDX was confirmed by analyzing the headspace of microcosms containing ring-labeled [15N]-RDX by GC-MS to monitor the masses at 45 Da (15N14NO). A Hewlett-Packard 6890 GC (Mississauga ON) coupled with a 5973 quadrupole mass spectrometer was used for this analysis. A GS-Gas Pro (30 m × 0.32 mm) capillary column (J & W Scientific, Folsom CA) was used under splitless condition. The column was maintained at -25 °C for 1.75 min, after which time it was raised to -10 °C/min for 3 min. Helium was the carrier gas, and the injector and detector were maintained at 150 and 280 °C, respectively. The injection volume was 15 µL. Liquid chromatography-mass spectrometry (LC-MS) was used to verify the presence of RDX and MNX. Analyte ionization was achieved in a negative electrospray ionization ES (-) mode producing mainly the deprotonated mass ions, [M - H]-. This system consisted of a Micromass Platform II benchtop single quadropole mass detector fronted by a Hewlett-Packard 1100 series HPLC system equipped with a photodiode array detector. Samples (50 µL) from the microcosms were injected into a Supelcosil LC-CN column (25 cm × 4.6 mm; 5 µm particle size) at 35 °C. The instrument conditions used are reported elsewhere (11). Solid-phase microextraction (SPME) in conjunction with GC-MS (HP 6890/MSD HP 5973 system) was used to detect formaldehyde (HCHO), a possible metabolite of RDX. An SPME fiber with poly(dimethylsoloxane)/vinylbenzene (Supelco, Oakville ON) was used with the derivatizing agent O-(2,3,4,5,6-pentafluorobenzyl)hydroxylamine hydrochloride (17 mg/mL) (25). The detection limit for formaldehyde was 100 µg/L. Methanol, another possible metabolite of RDX, was analyzed by a Perkin-Elmer Sigma 2000 GC with an AS 2000b autosampler (Norwalk, CT) and a HayeSep Q 80/100 column (6 ft × 0.125 in. o.d.) (Supelco, ON). The oven temperature was 115 °C, and the FID detector was at 250 °C. The carrier gas was N2 at 20 mL/min. The injection volume was 1 µL, and 2-propanol was used as an internal standard. The GC detection limit was 1 mg/L. Identity of methanol as an RDX metabolite was confirmed using [UL-14C]-RDX and by collecting the products (HPLC fractionation for methanol) for subsequent radioactivity measurement. The conditions of HPLC employed were the same as those that were just described for formic acid analysis. Retention time of methanol was determined using the same system used for formic acid analysis but with a refractive index detector. VOL. 34, NO. 16, 2000 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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FIGURE 2. RDX biodegradation by P. chrysosporium: (A) RDX disappearance and (B) MNX appearance with time (vertical bars denote standard error, 95% confidence). Aqueous samples were analyzed for NH4+ with an SP 8100 HPLC system equipped with a Waters 431 conductivity detector and a Hamilton PRP-X200 (250 mm × 4.1 mm × 10 µm) analytical cation exchange column using 30% methanol in 4 mM nitric acid at a flow rate of 0.75 mL/min. The injection volume was 100 µL. Analysis for NO2- and NO3- was performed by capillary electrophoresis using sodium borate (25 mM) and hexamethomium bromide (25 mM) as an electrolyte at pH 9.2 (23). The voltage was -20 kV, and the temperature was 25 °C. Samples were injected by applying 50 mbar pressure to the capillary inlet for 10 s. Detection was made at 215 nm, and the detection limit was 0.5 mg/L.
Results and Discussion RDX Disappearance/MNX Appearance. The fungus was able to reduce the quantity of RDX from 62 to 5 mg/L following 25 d of incubation (Figure 2A). Traces of the MNX metabolite appeared after 2 d and increased until 20 d, after which the MNX levels decreased (Figure 2B). Reddy and Gold (24) recently reported that reductive dechlorination reactions by cell-free enzyme systems were carried out by P. chrysosporium. Since nitroaromatic compounds (such as 2,4,6-trinitrotoluene) contain only nitro group subsituents, they are not direct substrates for the lignin-degrading enzymes (i.e., LiP and MnP) (25). Hence, P. chrysosporium reduces the aromatic nitro group to an amine in the first step in the degradation of nitroaromatic compounds (26). It is the amine products that are substrates for peroxidase-catalyzed oxidation, and the reduction reaction is the key step in the degradation of nitroaromatic compounds. Stahl and Aust (27) proposed that the plasma membrane redox system of P. chrysosporium reduced TNT to its two mono amino metabolites. Such a system may also explain the formation of MNX from RDX that was observed. To our knowledge, this is the first report on MNX formation from RDX biodegradation by P. chryrsosporium. The presence of MNX was confirmed by LC-MS, and the dinitroso and trinitroso metabolites of RDX (i.e., DNX and hexahydro-1,3,5-trinitroso-1,3,5-triazine (TNX), respectively) were never detected during the course 3386
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FIGURE 3. RDX biodegradation by P. chrysosporium: (A) [UL-14C]RDX mineralization to 14CO2, (B) appearance of N2O, and (C) MnP enzyme activity with time (vertical bars denote standard error, 95% confidence). of this study. For anaerobic degradation of RDX, McCormick et al. (34) postulated that if the nitroso group of MNX is reduced to a hydroxylamino group (hexahydro-1-hydroxylamino-3,5-dinitro-1,3,5-triazine), the molecule becomes unstable and hydrolytic ring cleavage occurs. In the present study, if the nitroso group of MNX was further reduced, there could have been a similar destabilization of the ring, as described by McCormick et al. (28). RDX Mineralization. Results of mineralization experiments show that 52.9% of RDX [UL-14C] was mineralized to 14CO after nearly 60 d (Figure 3A). In conjunction with this, 2 62.0% of the nitrogen from RDX was converted to nitrogen as N2O (Figure 3B). There was a 2-d lag period prior to detection of both 14CO2 and N2O. The MnP enzyme activity appeared after 2 d, and its level steadily increased until about 18 d and remained elevated until 30 d (Figure 3C). The evolution of N2O and CO2 correlate with MnP activity (Figure 3B,C). The LiP enzyme activity was absent in all microcosms. Experiments with ring-labeled [15N]-RDX, with subsequent analysis by GC-MS, confirmed that the N2O formed was N15N14O. From the mass spectra of 15N-labeled and unlabeled microcosms (Figure 4), it can be concluded that one nitrogen of N2O originated from the ring and the other from one of the nitro substituents of RDX. According to our calculations, the conversion of RDX to N2O represents about 62.0% of the nitrogen from the RDX. This is based on the fact that RDX was the main source of nitrogen in our microcosms and since N2O did not form in our chemical (RDX in medium with no fungus) or in our biological (fungus in medium with
TABLE 1. Summary of Carbon Balance for Microcosms Involving [UL-14C]-RDX and Nitrogen Balance for Unlabeled Microcosmsa source of 14C
% 14C recoveryb
source of N
% N recoveryb
2 in gas phase in aq phase of microcosms 14C recovered from biomass of fungus total 14C recovered
52.9 (( 0.7) 28.3 (( 4.8) 10.7 (( 5.7) 91.9 (( 11.2)
N2O in gas phase aq phase of microcosms recovered from biomass total N recovered
62.0 (( 3.1) nmc nm 62.0 (( 3.1)
14CO 14C
a
of
Both following 60 d of incubation (average values are reported, numbers in parentheses denote standard error, 95% confidence). C and N are percent of total from RDX. c nm, not measured.
b
Recovery
14
FIGURE 4. Mass spectra of (A) N2O produced in unlabeled microcosms and (B) 15N14NO in microcosms containing ring-labeled [15N]-RDX. no RDX) controls. Hence, the N2O originated from RDX, and it formed biotically from ring cleavage of RDX. Enzyme Activity. The MnP enzyme activity is depicted in Figure 3C for microorganisms containing RDX and the biological controls. The MnP enzyme may have reduced RDX to MNX (Figure 2b) since it has been found to catalyze reduction reactions in the presence of hydroquinones and Mn2+ (14). Since quinone (an oxidation product of hydroquinone) is present in all cells, hydroquinone may have acted with MnP to reduce RDX to MNX. Conversely, the MnP enzyme may have contributed to the oxidation of RDX. Scheibner and Hofrichter (29) studied the conversion of TNT and its reduction products (aminonitrotoluenes) by the MnP enzyme of the white rot fungus Nematoloma frowardii and the litter decaying fungus Stropharia rugosoannulata in cell free extracts. They found that MnP preparations from both fungi were able to mineralize amino-nitrotoluenes. The MnP enzyme generates Mn(III), a highly reactive intermediate, from Mn(II) (30). Mn(III) is stabilized by chelating organic acids (i.e., oxalate) produced by fungi (31). As well as being mobile, the chelated Mn(III) is a strong oxidizer, capable of one electron transfers. The fact that we did not observe RDX biodegradation products, aside from low levels of CH3OH (as discussed subsequently), is consistent with the observation that the one electron-transfer process of the lignin-degrading enzyme system gives rise to “enzy-
matic combustion” (32). Chemical studies have shown that, unlike nitroaromatic compounds, once the nonaromatic cyclic nitramines (i.e., RDX or the high melting explosive (HMX)) undergo a change in their molecular structure, the ring collapses to produce small nitrogen- (N2O, NO2, NH3) and carbon- (HCHO, HCOOH, and CO2) containing molecules (33). The cytochrome P450 enzyme system may have been involved in RDX metabolism. However, this enzyme system has not been extensively explored for P. chrysosporium (34, 35). LiP enzyme activity was not detected in any of the microcosms. We previously assayed LiP in this medium (36), hence its absence is not likely due to difficulty in its detection. However, LiP may have been a direct substrate for other peroxidases. Metabolites. Despite extensive efforts to identify metabolites or products from ring cleavage of RDX, CO2 and N2O were the predominant products observed in the present study. Intermediates may be short-lived due to the potent oxidizing capability of the MnP enzyme that was present in our system (37). The only other trace metabolite that was detected in our system was CH3OH. After 9 d incubation, 1.33 mg/L (( 0.04) CH3OH was measured in the aqueous phase of microcosms containing RDX. No CH3OH was detected in the biological controls. The presence of CH3OH as a metabolite of RDX was confirmed by HPLC fractionation of aqueous samples of [UL-14C]-RDX microcosms. Valli et al. (26) found that the MnP enzyme was capable of oxidizing 2-amino-4-nitrotoluene (2-AmNT) to 4-nitro-1,2-benzoquinone and CH3OH. As pointed out by Scheibner and Hofrichter (29), CH3OH formation demonstrates the ability of MnP to split off single-carbon fragments. To our knowledge, CO2 is the only product that has been identified from the biodegradation of RDX by P. chrysosporium (12, 13). Bose et al. (38) studied the byproducts from advanced oxidation of RDX in aqueous solution. They found that traces of 1,3-dinitro-1,3,5-triazacyclohex-5-ene formed from the removal of HNO2 from RDX during photolysis. If such a product formed in our experiments, it would have been a transient intermediate since the rate at which HNO2 is removed from RDX is 105 times slower than the rate at which the product (1,3-dinitro-1,3,5-triazacyclohex-5-ene) is cleaved (33). In our experiments, nitrite and nitrate were not detected in RDX microcosms, chemical controls, nor in the biological controls. Although there was an initial increase in NH4+ for the RDX microcosms and the biological controls (average of 32 mg/L at pH 4.5 for both microcosms), the levels fell below detection after 11 d. Carbon Balance. A considerable percentage of carbon from RDX remained in the aqueous phase (28.3 ( 4.8%, Table 1) after 60 d of incubation, and the structure of this moiety is unknown. Since there was a fraction of 14C recovered from the biomass (10.7 ( 5.7%), it is likely that a fraction of the nitrogen from RDX was also associated with the biomass. This is not surprising since RDX was a source of nitrogen. Although we did not complete a nitrogen balance for the biomass and aqueous phases of the microcosms (Table 1), a majority of nitrogen was liberated as N2O (62.0%). To the best of our knowledge, P. chrysosporium is not a known VOL. 34, NO. 16, 2000 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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nitrifier. Hence it is unlikely that N2O formation was from nitrification. Environmental Implications. That much of the nitrogen from RDX was converted to N2O (62.0%) may be relevant to monitoring natural attenuation of contaminated sites. As noted earlier, we identified rapid formation of N2O from anaerobic biodegradation of RDX (11). Although further research is necessary to determine the universality of RDX transformation to N2O, the potential utility of N2O monitoring as an indicator of RDX biodegradation is a notable possibility. There has been recent interest in the production of N2O by nitrifying and denitrifying bacteria because it is a greenhouse gas (39). Since there are a variety of microorganisms in the environment that produce and consume N2O (40), efforts have been made to monitor and quantify its sources in soil (39). For example, N2O measurements can be made with manual static chamber systems that are inexpensive, easy to perform, and nondestructive (39). Consequently, a site can be monitored for long periods while avoiding the problem of spatial heterogeneity of samples collected for sacrificial sampling. Since RDX is not extensively immobilized by soil (unpublished results), information concerning biotransformation products is important for monitored natural attenuation. Further research is focused on characterizing conditions of N2O formation from RDX in various soil and microbial systems.
(13) (14) (15)
(16)
(17) (18) (19) (20) (21) (22) (23) (24) (25)
Acknowledgments (26)
We thank the Natural Sciences and Engineering Research Council and the National Research Council (NRC) of Canada for a fellowship to T.W.S. and the Department of National Defence for their continued interest in this work. We also thank L. Paquet, A. Halasz, J. Hodgson, and R. Roy for insightful discussions and technical support. This is NRC Publication No. 43316.
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(32) (33)
(34) (35) (36) (37) (38) (39) (40)
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Received for review February 11, 2000. Revised manuscript received May 18, 2000. Accepted May 19, 2000. ES000998Y
