Metabolism, Microflora Effects, and Genotoxicity in Haloacetic Acid-Treated Cultures of Rat Cecal Microbiota

G. M. Nelson*,1, A. E. Swank*, L. R. Brooks*, K. C. Bailey{dagger} and S. E. George*

* U. S. Environmental Protection Agency, Office of Research and Development, National Health and Environmental Effects Research Laboratory, Environmental Carcinogenesis Division, Research Triangle Park, North Carolina 27711; and {dagger} Department of Biology, North Carolina Central University, Durham, North Carolina 27707

Received September 27, 2000; accepted December 19, 2000


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Haloacetic acids are by-products of drinking water disinfection. Several compounds in this class are genotoxic and have been identified as rodent hepatocarcinogens. Enzymes produced by the normal intestinal bacteria can transform some promutagens and procarcinogens to their biologically active forms. The present study was designed to investigate the influence of the cecal microbiota on the mutagenicity of haloacetic acids, and to look at changes in the microbiota populations and enzyme activities associated with exposure to haloacetic acids. PYG medium containing 1 mg/ml of monochloroacetic (MCA), monobromoacetic (MBA), dichloroacetic (DCA), dibromoacetic (DBA), trichloroacetic (TCA), tribromoacetic (TBA), or bromochloroacetic (BCA) acid was inoculated with rat cecal homogenate and incubated anaerobically at 37°C. Growth curves were performed with enumeration of the microflora populations on selective media. Mutagenicity in a Salmonella microsuspension bioassay was determined after incubation for various lengths of time, with or without the cecal microbiota. At 15 h of incubation, enzyme assays determined the activities for ß-glucuronidase, ß-galactosidase, ß-glucosidase, azoreductase, nitroreductase, dechlorinase, and dehydrochlorinase. The haloacetic acids, with the exception of BCA, were toxic to the cecal microbiota, and especially to the enterococci. DBA, TBA, and BCA were mutagenic in the microsuspension assay, but the presence of the intestinal flora did not significantly alter the mutagenicity. BCA increased the activities of several enzymes, and therefore has the potential to affect the biotransformation of co-exposed compounds.

Key Words: disinfection by-products; mutagenicity; Salmonella microsuspension assay; biotransformation; intestinal flora; enzymes..


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Chlorination of drinking water prevents the spread of waterborne infections and has been a common practice for almost a century. However, consumption of chlorinated drinking water has been associated with urinary and gastrointestinal tract cancers in human epidemiology studies (Koivusalo et al., 1994Go; Morris et al., 1992Go). Disinfection by-products (DBPs) arise from the reaction between natural organic material in the water and chlorine. A commonly occurring class of DBPs, the haloacetic acids, includes dichloroacetic acid (DCA) and trichloroacetic acid (TCA), which are hepatocarcinogenic in rodents (Bull et al., 1990Go; DeAngelo et al., 1991Go, 1996Go). DCA is mutagenic in the Salmonella reversion, microscreen prophage induction, and mouse lymphoma assays (DeMarini et al., 1994Go, Harrington-Brock et al., 1992Go), genotoxic in the SOS chromotest (Giller et al., 1997Go) and generates chromosomal aberrations (Fusco et al., 1996). DCA and TCA show mutagenic activity in Salmonella typhimurium strain TA100, with the Salmonella fluctuation test (Giller et al., 1997Go). The carcinogenicity of the bromo- and bromochloroacetates has not yet been evaluated. However, mono-, dibromo-, and tribromoacetic acids (MBA, DBA, and TBA) are mutagenic in strain TA100 in the Salmonella fluctuation assay, and DBA and TBA are genotoxic in the SOS chromotest (Giller et al., 1997Go). DBA or BCA, but not DCA or TCA, in the drinking water of B6C3F1 mice results in the formation of hepatic 8-hydroxydeoxyguanosine adducts (Parrish et al., 1996Go).

The enzyme activities of the normal mammalian intestinal microbial population can transform many promutagens and procarcinogens to their mutagenic and carcinogenic forms (Chadwick et al., 1992Go). For example, glycosides are compounds consisting of a non-sugar moiety (aglycone) bound to a sugar by an {alpha}- or ß-glycosidic linkage. Glycosides can enter the gut from two major sources, the diet or the liver (compounds detoxified by glucuronide formation in the liver are secreted into the intestine via the bile). The intestinal flora then can hydrolyze the ß-glucuronide bond, releasing the aglycones, some of which are toxic or carcinogenic (Goldin, 1986Go). Repeated exposure to a compound via the enterohepatic circulation can amplify its biological activity. The enzymes of the intestinal flora (specifically, ß-glucosidase) hydrolyze cycasin, a nongenotoxic plant glycoside, to a genotoxic aglycone, methylazomethanol (MAM; Brown and Dietrich, 1979). Cycasin causes tumors in rats with normal intestinal flora, but not in germ-free rats (Laquer et al., 1967Go). The principal glycosidases produced by the intestinal flora are ß-glucosidase, ß-galactosidase, and ß-glucuronidase. E. coli and Clostridium are associated with ß-glucuronidase activity. However, E. coli has low ß-glucosidase activity, with higher activities of this enzyme found with Bacteroides and Enterococcus faecalis (Hawksworth et al., 1971Go).

Many azo dyes are used in the food and textile industries. The bacterial flora can reductively hydrolyze the azo bond by the action of azoreductase, which results in the formation of substituted aromatic amines, a number of which are well established carcinogens. The reduction of nitro groups by microbial nitroreductase in the intestine can be another source of aromatic amines (Goldin, 1986Go). Both nitroreductase and ß-glucuronidase activity are necessary for 2,6-dinitrotoluene genotoxicity (George et al., 1994Go). Microbial dechlorinase activity may be involved in the metabolism of trichloroacetic and dichloroacetic acids in the gut (Moghaddam et al., 1996Go). The DBPs dichloro-, dibromo-, or bromochloroacetic acid, administered in the drinking water of Fischer 344 rats, impacted the intestinal metabolism and microflora populations, indicating the potential for altered bioactivation of co-administered promutagens and procarcinogens (George et al., 2000Go).

The objective of the current study is to determine whether in vitro cultures of the rat intestinal microbiota can metabolize monochloro-, dichloro-, trichloro-, monobromo-, dibromo-, tribromo-, and bromochloroacetic acids to intermediates that are more or less mutagenic than the parent compound. In addition, the effects of these DBPs on the intestinal microbial populations and their metabolism are studied, with emphasis on the enzymes often involved in the bioactivation of promutagens and procarcinogens.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Chemicals.
The disinfection by-products monochloroacetic acid (MCA, CAS# 79–11–8, 99+ %), dichloroacetic acid (DCA, CAS# 79–43–6, 99+%), trichloroacetic acid (TCA, CAS# 76–03–9, 99%), bromochloroacetic acid (BCA, CAS# 5589–96–8, 97%), monobromoacetic acid (MBA, CAS# 79–08–3, 99+%), dibromoacetic acid (DBA, CAS# 631–641–1, 97%), and tribromoacetic acid (TBA, CAS# 75–96–7, 99%) were purchased from Aldrich Chemical Company (Milwaukee, WI). The enzyme substrates p-nitrophenyl-ß-D-glucuronide (GLR, CAS# 137629–36–8), p-nitrophenyl-ß-D-galactopyranoside (GAL, CAS# 3150–24–1), and p-nitrophenyl-ß-D-glucopyranoside (GLU, CAS# 2492–87–7) were obtained from Sigma Chemical Company (St. Louis, MO). 3,4-Dichloronitrobenzene (DCNB, CAS # 99–54–7, 99%), 1,1-bis (4-chlorophenyl)-2, 2, 2-trichloroethane (p,p'-DDT, CAS# 502903, 98%), and methyl orange (CAS# 547–58–0, 85+%) were from Aldrich. Pre-reduced, anaerobically sterile (PRAS) peptone yeast glucose (PYG) media was purchased from Remel Microbiology Products (Lenexa, KS). Unless indicated below, all other chemicals were of reagent grade and obtained commercially.

Growth curves.
Forty-eight-h growth curves were performed in vitro with rat cecal homogenate and with each of 4 strains isolated from the rat cecum. Adult male CDF rats from Charles River Laboratory (Raleigh, NC) were provided food (Purina 5001, Purina Mills, St. Louis, MO) and water ad libitum. Rats were asphyxiated with CO2 and taken into an anaerobic chamber (Coy Laboratory Products, Inc., Grass Lake, MI) where the cecum was removed and placed into 20 ml of prereduced VPI buffer (2.0 g gelatin, 0.5 g cysteine, 500 ml deionized water, and 500 ml aqueous solution containing 0.1 g anhydrous CaCl2, 0.1 g anhydrous MgSO4, 0.5 g K2HPO4, 5.0 g NaHCO3, and 1.0 g NaCl) (Holdeman et al., 1977Go), and minced with scissors. Forty µl of this cecal homogenate was added to 10 ml of PYG media along with 0.1 ml of a 100 mg/ml aqueous solution of DBP (filter-sterilized) for a final concentration of 1 mg/ml. Tubes (2 per treatment) were vortexed and incubated anaerobically at 37°C. At 0, 3, 6, 12, 24, and 48 h of incubation, a 0.1 ml aliquot of culture was removed, diluted in prereduced phosphate-buffered saline, and plated onto selective media for enumeration of flora populations. Enumerated populations were: lactose-fermenting enteric bacilli on MacConkey agar (MAC), enterococci on KF streptococcus agar (KF), total anaerobes and facultatives on Brucella laked-blood agar (BA), anaerobic gram-negative bacilli on Brucella laked blood agar + vancomycin and kanamycin (VK), and lactobacilli on Rogosa agar (ROG). MAC and KF plates were incubated aerobically for 24 and 48 h, respectively. BA, VK, and ROG plates were incubated anaerobically for 72 h.

The 4 cecal isolates, chosen for growth curves because of their abundance, were identified as Bacteroides distasonis, Bacteroides uniformis, Clostridium bifermentens, and Lactobacillus johnsonii, using the methods and software of the Microbial Identification System (MIDI, Inc., Newark, DE). Overnight cultures (25 ml) of each strain were grown in PYG broth. Cultures were centrifuged, washed in 5 ml of reduced salts solution, then resuspended in 1 ml of reduced salts solution. To each 10 ml PYG tube was added 0.1 ml of a 100 mg/ml aqueous solution of DBP and 0.1 ml of bacterial suspension. Tubes (2 per treatment) were incubated and samples removed for dilution and plating (on BA only) as described previously. Growth curves for the isolates also were performed in salts solution (5 mg CaCl2 -3881µ2H2O, 4 mg anhydrous MgSO4, 20 mg K2HPO4, 20 mg KH2PO4, 200 mg NaHCO3, and 1.0 g NaCl in 1 liter deionized, distilled H2O, [Microbial Identification System operating manual, p. C-4, MIDI, Inc., Newark, DE]) instead of PYG, to determine if the isolates could maintain growth by utilizing the DBP as a carbon source.

Enzyme assays.
PYG tubes were inoculated in triplicate with a DBP (1 mg/ml) and with cecal homogenate or a cecal isolate, as described above for the growth curves. Because of its toxicity, MBA also was tested at a second concentration of 0.1 mg/ml. After 15 h of incubation, cultures were centrifuged, the pellet washed in 5 ml prereduced VPI buffer, and resuspended in 10 ml VPI buffer. A 1 ml aliquot of each sample was transferred into 9 ml of 0.85% saline for dilution and plating (enumeration on BA only), and for APIZYM (bioMerieux Vitek, Inc., Hazelwood, MO) enzyme analysis following the manufacturer's instructions. The experiment was repeated twice for each DBP.

For determination of GLR, GAL, and GLU activities, 2 ml aliquots of each sample (pellet suspended in VPI buffer, see above paragraph) were transferred to 3 separate tubes, each containing 8 ml of VPI buffer. A rubber septum cap was placed on each tube, and tubes were removed from the chamber and placed on ice. A 37.5 µl aliquot of DMSO (culture controls and reagent controls) or the substrate mix containing 170 mg/ml of the nitroreductase competitor 3,4-dichloronitrobenzene and 130 mg/ml of the specific substrate GLR, GAL, or GLU was injected into each tube and incubated at 37°C with shaking for 1 h (Chadwick et al., 1995Go). Placing the tubes on ice terminated the reactions. Particulate matter was sedimented by centrifugation at 4°C. Tubes were again placed on ice and the OD was recorded at 405 nm using a Spectronic 20+ (Spectronic Instruments, Inc., Rochester, NY). The concentration of released p-nitrophenol was calculated from a p-nitrophenol standard curve.

For determination of nitroreductase, azoreductase, dechlorinase, and dehydrochlorinase activities, septum caps were placed on the tubes containing the remaining 3 ml of sample. The tubes were removed from the chamber and placed on ice. Twenty-five µl of DMSO only (culture controls and reagent controls) or 25 µl of substrate mix containing 25 mg/ml of 3,4-dichloronitrobenzene, 12.5 mg/ml of p,p'-DDT, and 23 mg/ml of methyl orange was injected through the septum cap of each tube. Tubes were incubated for 1 h at 37°C, then placed on ice to terminate the reaction. Samples were extracted, derivatized, and analyzed by gas chromatography according to the method of Chadwick et al. (1993).

Salmonella microsuspension bioassay.
A microsuspension modification of the Ames Salmonella reversion assay (DeMarini et al., 1989Go) was used to compare the mutagenicity of the DBP compounds/metabolites after incubation for various lengths of time, with or without the cecal microbiota. The microsuspension assay was chosen because of its greater sensitivity with a smaller mass of sample than that required by the standard plate incorporation assay. Growth curves in PYG were performed as previously described, with unopened sample tubes (2 per treatment) frozen (–20°C) at 0, 3, 6, 12, and 24 h of incubation. The concentration of haloacetic acid in the PYG cultures was 1 mg/ml, with the exception of MBA, which was at 0.1 mg/ml. Samples were extracted with methyl tert-butyl ether (MTBE), blown to dryness, and dissolved in 100 µl DMSO, then frozen at –20°C until bioassay.

Based on the results of previous mutagenicity testing of these compounds in Salmonella (data not shown), tester strain TA100 was used for bioassay of the brominated compounds and strain TA104 was used for the chlorinated compounds. BCA was tested with strain TA100. The tester strains were grown overnight in Oxoid Nutrient Broth No. 2 and harvested by centrifugation. The pellet was resuspended in 0.015 M phosphate buffer (pH 7.4) at either a 5x (TA100) or a 1x (TA104, spontaneous counts were too high with a 5x culture) concentration of the overnight culture. To each bioassay tube (13 x 100 mm, glass disposable), we added 5 µl of the sample or sample dilution (1:2.5 dilution), 100 µl of 0.015 M phosphate buffer (without S9) or 100 µl S9 mix (3% v/v S9 concentration), and 100 µl of cells. Each sample was run in duplicate. Tube racks were covered with foil and incubated at 37°C without shaking for 90 min, then placed on ice. Molten top agar (2.5 ml) was added to each tube, which was vortexed and poured onto a VBME plate. Plates were incubated for 72 h at 37°C and counted on an Artex 880 colony counter. DMSO was used as the negative control. Sodium azide (1 µg/plate) was the positive control (–S9) for strain TA100 and methyl glyoxal (25 µg/plate) for strain TA104. 2-Aminoanthracene (0.25 µg/plate) was the positive control (+S9) for TA100; 2-aminoanthracene (2.5 µg/plate) was the positive control for TA104. Assuming 100% recovery of the DBP after extraction, the dose of DBP would be 500 µg/plate, or 200 µg/plate for diluted samples (50 and 20 µg/plate for MBA). Actual recoveries (averages) are reported in Table 1Go.


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TABLE 1 Recovery of Disinfection By-products after Extraction (Salmonella Microsuspension Bioassay)
 
Statistical analyses.
All statistical analyses were done using SigmaStat 2.0 (SPSS Science, Inc., Chicago, IL). Effect of the haloacetic acids on the cecal microbiota populations was analyzed using two-way analysis of variance (ANOVA, DBP treatment x time, n = 2) with Dunnett's multiple comparison method for all means vs. the control. For enzyme analysis, one-way analyses of variance were performed (n = 6 for quantitative results, n = 2 for APIZYM results) with Dunnett's comparison of all means vs. the control. A positive mutagenic response was reported only when a 2-fold or greater increase of revertants over the spontaneous level was present. Mutagenicity also was analyzed by two-way ANOVA (treatment = chemical only, chemical + cecal homogenate, or cecal homogenate only x time; n = 2) with Bonferroni's all pairwise multiple comparison procedure, to determine if the mutagenicity of the haloacetic acids was altered by the cecal microbiota. A significance level of p <= 0.05 was used throughout.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Growth Curves
Cultures of rat cecal microbiota exhibited the typical growth curve for bacterial populations, including a period of exponential growth followed by a stationary phase. In the interest of conserving space, individual growth curves are not shown. However, Table 2Go reports values averaged over all time points for the enumerated populations, and its examination leads to the same conclusions as a visual inspection of the growth curves. The haloacetic acids differed in their toxicity to the cecal microbiota. Generally, the trihalogenated acetic acids and bromochloroacetic acid least affected growth, whereas the monohalogenated acetic acids were the most toxic (Table 2Go). Monobromoacetic acid was particularly toxic. Microbial numbers from all 5 media types were significantly lower for cultures containing monobromoacetic acid than for control cultures. Bromochloroacetic acid was the only haloacetic acid that did not exhibit toxicity toward any of the enumerated microbial populations. The enterococci population was the most sensitive to the toxic effects of haloacetic acids, with all but TBA and BCA resulting in significant reductions (Table 2Go).


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TABLE 2 Enumeration of Cecal Microbial Populations on Selective Media
 
Similar results for toxicity were obtained for pure culture growth curves with the selected cecal isolates (data not shown). Monobromoacetic acid was by far the most toxic of the compounds studied. Monochloroacetic acid and dibromoacetic acid also were very toxic to the Bacteroides strains. Growth did not occur in the minimal salts medium, indicating that the isolates were unable to use haloacetic acids as sole carbon and energy source.

Quantitative Enzyme Assays
In cultures of rat cecal flora, the highest activity of the 7 enzymes measured (Fig. 1Go) was for ß-galactosidase, with the exception of the BCA treatment group where ß-glucosidase levels were very high (significantly higher than in the control group). In addition, dehydrochlorinase activity was significantly higher in the BCA treatment group than in the control group (Fig. 1Go, Table 3Go).



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FIG. 1. Enzyme activities reported as µg metabolite/106 CFU/h. Metabolites are 3,4-dichloroaniline (nitroreductase), p-nitrophenol (ß-glycosidases), N,N-dimethyl-p-phenylenediamine (azoreductase), p,p'-DDE (dehydrochlorinase), and p,p'-DDD (dechlorinase).

 

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TABLE 3 Summary of Significant in Vitro and in Vivo Changes in Microflora Enzyme Activities following DBP Treatment
 
Enzyme activities in pure cultures of rat intestinal isolates were strain dependent (Fig. 1Go). Very little enzyme activity (less than 2 µg substrate/106 CFU/h) was detected in cultures of C. bifermentens or L. johnsonii for any treatment group (data not shown). ß-Glycosidase activity, particularly ß-galactosidase activity, was present in Bacteroides cultures. Compared to the control culture, ß-galactosidase activity was significantly decreased in cultures of B. distasonis and B. uniformis for all treatment groups. This decrease was not seen in cultures of rat cecal flora. In cultures of B. uniformis, ß-glucosidase activity was significantly lower than the control group for all treatment groups except BCA, where, as with the rat cecal flora, the activity was significantly higher (Fig. 1Go, Table 3Go).

Semiquantitative Enzyme Assays
Semiquantitative analysis of 19 enzymes was performed using APIZYM assays (Fig. 2Go). Table 3Go reports only those API results where statistical significance is shown. The following 11 enzymes were detected in control cultures of rat cecal flora: alkaline phosphatase, esterase (C4), esterase lipase (C8), leucine arylamidase, acid phosphatase, naphthol-AS-BI-phosphohydrolase, {alpha}-galactosidase, ß-galactosidase, {alpha}-glucosidase, ß-glucosidase, and N-acetyl-ß-glucosaminidase. Interestingly, ß-glucuronidase was not detected in control cultures but was detected in MCA-, MBA-, and DBA-treated cultures. These cultures also had the highest levels of ß-glucuronidase in the quantitative assays, but significance was not demonstrated. In most cases the API results correlated well with the quantitative enzyme results. However, because of the semiquantitative nature of the assay and the small sample sizes, statistical significance is difficult to demonstrate. Quantitative assays showed an elevated level of ß-glucosidase for BCA-treated cultures, and this effect was also seen using APIZYM, although it was not statistically significant. ß-Galactosidase activity was significantly reduced in cultures treated with TBA or MBA. Quantitative assays also showed a reduction in ß-galactosidase activity for these 2 treatment groups, but statistical significance was not shown. For the enzyme leucine arylamidase, activity was significantly lower in MCA-, MBA-, and DBA-treated cultures than in the control. Acid phosphatase activity was significantly lower in MCA- and MBA-treated cultures. Alkaline phosphatase activity was significantly elevated in BCA-treated cultures.



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FIG. 2. APIZYM activities. Enzyme numbers are not shown on graphs for enzymes with no activity. 1, control; 2; phosphatase alkaline; 3, esterase; 4, esterase lipase; 5, lipase; 6, leucine arylamidase; 7, valine arylamidase; 8, cystine arylamidase; 9, trypsin; 10, chymotrypsin; 11, phosphatase acid; 12, napthol-AS-BI-phosphohydrolase; 13, {alpha}-galactosidase; 14, ß-galactosidase; 15, ß-glucuronidase; 16, {alpha}-glucosidase; 17, ß-glucosidase; 18, N-acetyl-ß-glucosaminidase; 19, {alpha}-mannosidase; and 20, {alpha}-fucosidase.

 
As expected, fewer enzymes demonstrated activity in the pure cultures (Fig. 2Go). Again, very little enzyme activity was detected in cultures of C. bifermentens or L. johnsonii (data not shown). For B. distasonis, the active enzymes were alkaline phosphatase, esterase lipase, leucine arylamidase, acid phosphatase, naphthol-AS-BI-phosphohydrolase, {alpha}-galactosidase, ß-galactosidase, {alpha}-glucosidase, and N-acetyl-ß-glucosaminidase. As with the quantitative assays, ß-galactosidase activity was suppressed relative to the control in MCA-, DCA-, MBA-, DBA-, and TBA-treated cultures. Alkaline phosphatase activity was significantly depressed by all treatments. N-acetyl-ß-glucosaminidase activity was significantly reduced by all treatments except DCA and TCA. Acid phosphatase activity was significantly reduced in all treatment groups. For B. uniformis, the active enzymes were alkaline phosphatase, leucine arylamidase, acid phosphatase, naphthol-AS-BI-phosphohydrolase, and ß-galactosidase. Although statistical significance was not shown, APIZYM assays detected decreased ß-galactosidase activity for all treatment groups, mirroring the results of the quantitative assay. Both Bacteroides strains treated with the lower dose of MBA showed extremely high activity for the enzyme naphthol-AS-BI-phosphohydrolase. However, statistical significance could not be shown because of the high variability.

Salmonella Microsuspension Bioassays
No mutagenicity was detected for MCA, DCA, or TCA with S. typhimurium strain TA104, therefore these data are not shown. Mutagenicity (>= 2-fold increase over spontaneous) was observed in TA100 for samples containing DBA or BCA with and without S9. TBA was mutagenic only in the presence of S9. Positive control values for TA100 (average of 4 experiments) were sodium azide (–S9, 449 revertants/plate) and 2-aminoanthracene (+S9, 909 revertants/plate).

Undiluted samples containing cecal homogenate often were toxic to Salmonella when S9 was not present, especially at the later time points. Because toxicity was present without S9 for undiluted samples containing DBA and cecal homogenate, only the results for the diluted samples were plotted (Fig. 3aGo). Samples containing DBA only were mutagenic over the entire time course. Mutagenicity of the samples containing cecal homogenate only and samples containing DBA+ cecal homogenate (cec) dropped sharply by 6 h of incubation. With S9 added, the undiluted samples were more mutagenic than the diluted samples (Fig. 3bGo). Samples containing DBA only were significantly more mutagenic than those containing cecal homogenate only, and mutagenicity for samples containing both the chemical and cecal homogenate were intermediate between the two, but not significantly different from either. This was the only sample set for which samples containing cecal homogenate alone were mutagenic.



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FIG. 3. Mutagenicity of haloacetic acids in microsuspension bioassay with Salmonella typhimurium strain TA100; chemical only (filled circle), chemical + cecal homogenate (filled square), cecal homogenate only (*). Solid line denotes undiluted samples; dashed line denotes 1:2.5 sample dilution. A solid line with no marker indicates the spontaneous level.

 
Undiluted samples containing TBA were toxic to S. typhimurium TA100 without S9, but were slightly mutagenic in the presence of S9 (Fig. 3cGo). Revertants/plate were significantly higher in TBA and TBA + cec samples than in samples containing cecal homogenate only. The TBA and TBA + cec samples did not differ significantly. Results with the diluted samples were similar, and therefore are not graphed.

The undiluted samples were more mutagenic than their dilutions for samples containing BCA (Figs. 3d and 3eGo). This was the only sample set with strain TA100 for which undiluted samples containing cecal homogenate were not toxic at 12 and 24 h. Without S9, undiluted samples containing BCA only or BCA + cec were mutagenic and significantly different from samples containing cecal homogenate only, but not from each other. The diluted samples were not mutagenic. With S9, both diluted and undiluted samples containing BCA only or BCA + cec were mutagenic and significantly different from the samples containing cecal homogenate only. BCA and BCA + cec samples differed significantly from each other, only for undiluted samples, at the 3 h time point, when the revertants/plate were significantly higher in samples containing both BCA and cecal homogenate. This difference was not present in the diluted samples.


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Concentrations of haloacetic acids in finished drinking water are dependent on the source water and the disinfection method. The U.S. Environmental Protection Agency regulates disinfection by-products in drinking water and has set the maximum contaminant level (MCL) for total haloacetic acids (MCA, MBA, DCA, DBA, and TCA) at 60 ppb (U.S. EPA, 1998Go). The haloacetic acid concentrations used in the present study (1 mg/ml) were significantly higher than those found in finished drinking water. At this concentration, all of the haloacetic acids, except for BCA, were toxic to at least some population of the rat cecal microbiota. Enterococci appear to be particularly sensitive to haloacetic acids. In Fisher 344 rats exposed to this same concentration (1 g/l) of DCA, DBA, or BCA in the drinking water, DCA and DBA (but not BCA) were toxic to enterococci in the small and large intestine, respectively, but cecal populations remained constant (George et al., 2000Go). In the current in vitro study, DCA and DBA, as well as MCA, TCA, and MBA (but not TBA or BCA) were toxic to the cecal enterococci of CDF rats. Whether chronic exposure to more realistic concentrations of haloacetic acids would affect the microbiota is not known. However, disturbing the microbial balance of the intestinal tract can result in increased susceptibility to invading pathogens by reducing the resistance to colonization that is normally provided by the indigenous flora, known as "colonization resistance" or "competitive exclusion" (Van der Waaij, 1971, 1989Go). Due to the complexity of the intestinal flora, it is often difficult to show subtle changes in the abundance of specific bacterial populations. However, changes in the metabolic activity of the flora as a whole may be easier to measure and of more relevance to the host.

Interestingly, the only chemical that was not toxic to the rat cecal microbiota (BCA) was also the only chemical to significantly alter the enzyme activity of the microbiota in vitro, resulting in increased ß-glucosidase and dehydrochlorinase activities. Induction of dehydrochlorinase activity could be due to the availability of the haloacetic acid for metabolism. Enterococcus faecalis and Bacteroides spp. have been associated with ß-glucosidase activity (Hawksworth et al., 1971Go). While the enterococci population in vitro was reduced by treatment with most of the haloacetic acids, enterococci numbers were not affected by BCA treatment, and ß-glucosidase activity increased. Evidence for the contribution of Bacteroides to the increase in ß-glucosidase activity with BCA treatment was seen in pure cultures of B. uniformis. In vivo, ß-glucosidase activity was increased in the large intestine, but not in the cecum, by DCA, DBA, and BCA (George et al., 2000Go). Elevated enzyme activities indicate increased potential for bioactivation of promutagens and procarcinogens.

Azoreductase, nitroreductase, and dechlorinase activities were significantly reduced by one or more treatments in vivo, but not in vitro. Table 3Go summarizes in vitro and previous in vivo (George et al., 2000Go) week 5 results, indicating statistically significant changes in enzyme activity. DBP effects on enzyme activity in vivo often varied over the course of 5 weeks, making in vitro/in vivo comparisons difficult.

In cultures of B. distasonis and B. uniformis, but not the rat cecal cultures, ß-galactosidase activity was reduced for all treatment groups. In vivo, DCA, DBA, and BCA resulted in reduced cecal ß-galactosidase activity at weeks 1 and 3. However, by week 5, ß-galactosidase activity had returned to control levels in the DCA and DBA treatment groups, and had increased above control levels in the BCA treatment group.

APIZYM results correlated well with the quantitative results for those enzymes analyzed by both methods. Trends were similar, even though statistical significance was not always demonstrated. The haloacetic acids usually lowered the activities of additional enzymes assayed by the APIZYM methods. These enzymes are not known to be involved in the biotransformation of xenobiotics. In general, reduced enzyme activities may indicate a depressed cell metabolism, which in this case could be attributed to chemical toxicity. The lone exception again occurred with BCA-treated cultures of rat cecal flora where alkaline phosphatase activity was significantly elevated. BCA was not toxic to the rat cecal microbiota.

DBA, TBA, and BCA were mutagenic in TA100. After 3 h of incubation with BCA, when the culture of rat cecal flora was in exponential growth and should be most enzymatically active, a metabolite may have formed which was more mutagenic than BCA itself. However, overall, the co-incubation of the cecal microbiota with haloacetic acid either did not affect the mutagenicity of the chemical or reduced it.

In one sample set (with cecal microbiota derived from one animal) the cecal homogenate itself was mutagenic. Fecal mutagenicity has been well documented and is reviewed by Goldin (1986). The majority of fecal mutagens are fecapentaenes. A precursor, which is either a product of other bacteria in the colon, a result of diet, or a metabolite derived from the host, is necessary for fecapentaene production, in combination with Bacteroides. Fecal mutagenicity appears to be dependent on diet; humans on a typical "western" diet are at higher risk for colon cancer and have higher fecal mutagenicity than those on a vegetarian diet. However, all the rats in this study were on the same diet and housed identically. Therefore, individual genetic variability, or a differing flora composition due to a previous exposure are the most likely explanations for the occurrence of mutagenic cecal contents in only one animal.

Undiluted samples containing cecal homogenate often were toxic to Salmonella in the microsuspension assay. This was especially notable without S9 for the 12 and 24 h time points. By 12 h the cecal cultures were in stationary phase, during which growth is limited by the depletion of nutrients and the accumulation of toxic waste products in the medium. The addition of mammalian liver enzymes in the S9 mix resulted in detoxification of these samples.

In summary, high levels of haloacetic acid in the culture medium, with the exception of BCA, are toxic to the rat cecal microbiota, especially the enterococci. This effect also was seen in vivo in the intestine, and could increase the animals' susceptibility to disease. The mutagenicity of the haloacetic acids is not a result of biotransformation by the intestinal flora, but factors could be involved in the animal that cannot be mimicked in vitro (enterohepatic circulation, for example). BCA did elevate the activity of several enzymes involved in the biotransformation of xenobiotics, an effect that could potentially increase the biological activity of co-administered compounds. Therefore, future work should investigate this potential with compounds routinely found in drinking water.


    ACKNOWLEDGMENTS
 
The authors would like to thank Ms. Sarah Warren and Ms. Joanne Callahan for their valuable technical assistance, and Drs. Larry Claxton and Gary Held for their critical review of this manuscript.


    NOTES
 
This manuscript has been reviewed by the National Health and Environmental Effects Research Laboratory, U.S. Environmental Protection Agency, and approved for publication. Approval does not signify that the contents necessarily reflect the views and policies of the Agency, nor does mention of trade names or commercial products constitute endorsement or recommendation for use.

1 To whom correspondence should be addressed at U.S. EPA, 86 TW Alexander Drive, MD 68, Research Triangle Park, NC 27711. Fax: (919) 541-3966. E-mail: nelson.gail{at}epa.gov. Back


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