- Email: [email protected]
Growth of a bacterial consortium on triclosan
FEMS Microbiology Ecology 36 (2001) 105^112
www.fems-microbiology.org
Growth of a bacterial consortium on triclosan Anthony G. Hay a
a;b;c
, Peter M. Dees b , Gary S. Sayler
a;
*
Center for Environmental Biotechnology, 676 Dabney Hall, University of Tennessee, Knoxville, TN 37996-1605, USA b Department of Microbiology, Cornell University, Ithaca, NY 14853, USA c Institute for Comparative and Environmental Toxicology, Cornell University, Ithaca, NY 14853, USA Received 26 September 2000; received in revised form 6 April 2001; accepted 8 April 2001 First published online 21 May 2001
Abstract Triclosan is a polychlorinated hydroxy diphenylether that has been widely used as an antimicrobial compound. An enrichment using triclosan as a sole source of carbon and energy yielded a consortium of bacteria capable of growing on this compound. The dichloro ring was partially mineralized, resulting in the conversion of approximately 35% of the [14 C]triclosan to [14 C]CO2 . Use of molecular fingerprinting techniques and 16S rDNA cloning and sequencing aided in the identification and eventual isolation of an auxotrophic Sphingomonas-like organism, strain Rd1, which was able to partially mineralize triclosan when grown on complex media. ß 2001 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. Keywords : Biodegradation; Consortium ; Sphingomonas ; Triclosan; Antimicrobial
1. Introduction Triclosan is a broad spectrum antimicrobial compound that has recently found increased use in personal care products such as soaps, handcreams, toothpastes, and acne creams and in commercial products such as hosiery and plastics [1,2]. Evidence suggests that triclosan blocks fatty acid synthesis by interacting with the NADP binding site of the enoyl-ACP reductase, FabI in Escherichia coli [1] and InhA in Mycobacterium tuberculosis [3]. However, Streptococcus pneumoniae, which is also sensitive to triclosan, does not contain a FabI homolog [4]. Instead S. pneumoniae contains FabK, an isofunctional FAD-dependent enoyl-ACP reductase which is not sensitive to triclosan [4]. This suggests that triclosan must have another target in S. pneumoniae [4]. Pseudomonas aeruginosa, which is highly resistant to triclosan, contains both FabI and FabK, in addition to a multidrug e¥ux pump which is able to transport triclosan out of the cell [5]. Triclosan, otherwise known as Irgasan, is the selective ingredient in Pseudomonas Isolation Agar (PIA) [6].
* Corresponding author. Tel. : +1 (423) 974-8080; Fax: +1 (423) 974-8086; E-mail: [email protected]
Early work done by Voets et al. [7] demonstrated that up to 50% of the triclosan added to synthetic sewage sludge was degraded under both aerobic and anaerobic conditions via cometabolism. In minimal media, however, no degradation was observed. The authors therefore suggested that triclosan might persist in oligotrophic waters if it had not ¢rst been subjected to e¤cient wastewater treatment procedures [7]. Although there is little evidence to suggest widespread environmental accumulation or recalcitrance, several studies have demonstrated the presence of triclosan in sediments [8,9] and biota [10,11]. Concern about the environmental toxicity of the compound comes in part from the predioxin nature of triclosan which, upon exposure to heat or radiation, can undergo ring closure to form chlorinated dioxins [10]. This concern is compounded by the fact that di- and trichlorinated dibenzodioxins have been shown to contaminate commercial triclosan preparations [12]. Given concern about the widespread use of antimicrobial compounds and the possible emergence of drug-resistant bacteria, we sought to assess the potential for biodegradation of triclosan by bacteria in activated sludge from a residential wastewater treatment facility. In this study we describe the isolation and phylogenetic characterization of six strains from a consortium of bacteria able to grow on triclosan as a sole source of carbon and energy.
0168-6496 / 01 / $20.00 ß 2001 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. PII: S 0 1 6 8 - 6 4 9 6 ( 0 1 ) 0 0 1 2 7 - 1
FEMSEC 1245 5-7-01
106
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
2. Materials and methods 2.1. Bacteriological media Enrichments and subsequent cultivation of the consortium were done in a de¢ned, chloride-free minimal salts medium (MSM) containing 10 mM K2 HPO4 , 3 mM NaHPO4 , 10 mM (NH4 )2 SO4 , and 1 mM MgSO4 in deionized water. A chloride-free trace elements solution was also added at 1 ml l31 [13]. Consortium members were also streaked on MSM plates containing 1000 mg l31 sodium acetate (MSMA), 500 mg l31 triclosan as a sole source of carbon and energy (MSMT), with and without 50 mg l31 yeast extract (MSMTYE) and on the following complex media: Luria^Bertani medium (LB), PIA (Difco, Detroit, MI, USA) and YEPG. YEPG consists of 0.2 g of yeast extract, 2 g of peptone, 0.2 g of ammonium nitrate, and 1 g of glucose in 1 l of distilled water [14]. All solid media were solidi¢ed with 15 g l31 bacteriological agar or noble agar in the case of the minimal media. 2.2. Reagents Triclosan (5-chloro-2-(2,4-dichlorophenoxy)phenol) and [ C]triclosan were a gift from Ciba Specialty Chemical (Basel, Switzerland) and were 99% and s 99% pure respectively. The [14 C]triclosan had a speci¢c activity of 5.2 MBq mg31 and was uniformly labeled in the dichloro ring, while the other ring was unlabeled. Reagent grade ethyl ether, acetonitrile and Scintisafe Econo 1 scintillation cocktail were obtained from Fisher Scienti¢c (Pittsburgh, PA, USA). N-Nitrosyl-N-methylurea, which was used to generate diazomethane [14], was obtained from Sigma Chemical Company (St. Louis, MO, USA). 14
2.3. Bacterial cultures and conditions Activated sludge from the Neyland Wastewater treatment plant (Knoxville, TN, USA) was used as a source of inoculum for enrichments on triclosan. Initially, 500 mg of powdered triclosan was added directly to 1 l of activated sludge and incubated for 1 month at 28³C while being shaken at 150 rpm. After the initial 1 month incubation and subsequently every 2 weeks thereafter, the enrichment culture was transferred (10% v/v) to fresh MSM containing 500 mg l31 triclosan. Two weeks after the ¢fth transfer, the enrichment was streaked onto all of the media listed above and plates were incubated at 28³C. Individual colonies with distinctive morphologies were picked from the complex media plates and streaked on the same medium for single colony isolation, then restreaked on MSMT and MSMTYE plates. The enrichment culture was serially diluted and plated on MSMT and MSMTYE agar plates, which were then incubated for 1^3 weeks at room temperature. After incubation, the plates were wiped with a presoaked sterile cot-
ton swab, resuspended in 100 ml MSMT, incubated at 28³C and examined for an increase in visible turbidity [15]. In addition, to assess the possibility that numerically inferior organisms were required for triclosan mineralization, aliquots amounting to 10 ml of enrichment culture were centrifuged at 8000Ug in a microcentrifuge, the supernatant was decanted and the combined cell pellets were resuspended in 0.1 ml 30 mM phosphate bu¡er (pH 7.3). The 100U concentrated cell suspension was spread on MSMT and MSMTYE plates and incubated at 28³C for as long as 1 month. The plates were then wiped as above and resuspended in MSMT to check for growth. When grown in MSMT broth, the consortium often clumped. To avoid variation when optical density was used as a measure of growth, the growth of the consortium was monitored as an increase in protein content. The enrichment culture was vigorously shaken before sampling to disperse clumped cells and after dispersal 250-Wl aliquots of culture were transferred to individual 1.5-ml microcentrifuge tubes and frozen until the end of the experiment. Frozen cells were thawed, then permeabilized by adding 5 Wl of toluene and the protein concentration was measured using the Bio-Rad Protein Assay kit (Bio-Rad, Richmond, CA, USA) according to the manufacturer's instructions. 2.4. Mineralization assay and metabolite analysis The consortium and individual isolates were screened for triclosan mineralization by placing 5 ml of a triclosan-grown (consortium) or YEPG-grown, exponential phase (isolates) culture in a 40-ml EPA vial containing a smaller 8-ml glass vial to which 1 ml of 1 M NaOH was added. A mixture of unlabeled and 14 C-labeled triclosan in acetone was added to the culture to give a ¢nal concentration of 500 mg l31 and 250 000 dpm. The 40-ml EPA vials were then capped and shaken at 150 rpm while incubating at 28³C. The full volume of spent NaOH was periodically withdrawn and replaced with a fresh 1.0 ml of 1 M NaOH. The spent NaOH was added to 10 ml scintillation cocktail and [14 C]CO2 accumulation was measured using a Beckman LS5000CE liquid scintillation counter (Beckman, Fullerton, CA, USA). No growth was observed when acetone was added as a sole source of carbon and energy. After the cells were pelleted by microcentrifugation, the consortium supernatant was analyzed for the accumulation of metabolites by high pressure liquid chromatography (HPLC) using a Supelcosil LC-18-T reverse phase C18 column (Supelco, Bellefonte, PA, USA) attached to a binary LC 250 series pump with a model LC-235 diode array detector (both Perkin Elmer, Foster City, CA, USA). Absorbance was monitored at 210 nm. The eluent was 50% 40 mM acetic acid and 50% acetonitrile. Samples of the culture supernatant were also analyzed by gas chro-
FEMSEC 1245 5-7-01
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
107
matography. After centrifuging 100 ml of culture (10 000Ug for 10 min), the supernatant was acidi¢ed with HCl and then extracted three times with ethyl ether. The extracts were pooled, evaporated to dryness, then resuspended in 1.0 ml of ethyl ether and either derivatized with diazomethane [16] or analyzed in the underivatized state. Analysis of extracts was performed on a Hewlett Packard 6890 series gas chromatograph equipped with an HP5ms column (30 mU250 WmU0.25 Wm) and a 5973 series mass selective detector (GC-MS) (Hewlett Packard, Wilmington, DE, USA). The injector was held at 250³C. The initial oven temperature was 40³C and was ramped after 1 min at 10³C min31 to 250³C at which point the temperature was held constant for 10 min.
conditions were based on a modi¢ed touchdown protocol [22]. The initial denaturation step for the ¢rst primer set was 5 min at 95³C followed by 10 cycles of denaturation at 95³C for 45 s, annealing at 65³C for 15 s, and extension at 70³C for 60 s. For touchdown PCR, the annealing temperature was decreased by 1³C cycle31 for the ¢rst 10 cycles, followed by 20 cycles of annealing at 55³C. A 3-min extension at 70³C was included after completion of the 30 cycles of touchdown ampli¢cation. PCR products ampli¢ed from the ribosomal intergenic spacer region were separated in a 2% agarose gel after electrophoresis for 4 h at 80 mV.
2.5. Chloride determination
PCR was also used to amplify nearly full-length sections of the 16S rRNA gene from the consortium DNA lysate and individual isolates using universal primers 27F (AGAGTTTGATCMTGGCTCAG) and 1492R (TACGGYTACCTTGTTACGACTT). Touchdown thermal cycling conditions for the 27F/1492R primer set were similar to the protocol for RISA PCR except that the initial annealing temperatures for the ¢rst 10 cycles decreased at 1³C cycle31 from 55³C to 45³C, followed by 20 cycles of annealing at 45³C. PCR products ampli¢ed with the 27F/1492R primer pair were ligated into pCR2.1-TOPO (Invitrogen, Carlsbad, CA, USA) following the manufacturer's instructions. The ligation mixture was electroporated into DH5K electrocompetent cells and plated onto LB containing 50 mg l31 kanamycin and 40 mg ml31 X-gal. Plasmids were puri¢ed from white transformant colonies using Mini Prep Express Matrix (Bio 101, Carlsbad, CA, USA) following the manufacturer's instructions and plasmid DNA was screened for the presence of a 1500-bp insert after digestion with EcoRI and electrophoresis of the plasmid DNA in a 1% agarose gel. After screening for inserts, plasmids from 12 of the transformants were used for sequencing templates at the University of Tennessee, Knoxville (UTK) Sequencing Facility using primer 1492R. PCR products ampli¢ed directly from the consortium DNA extract and from individual isolates using the 27F/ 1492R primer set were digested with HhaI and separated in a 4% agarose gel after electrophoresis for 12 h at 25 mV. The rRNA genes from ¢ve isolates which gave either a unique BOX, RISA, or ampli¢ed ribosomal DNA restriction analysis (ARDRA) [23]) pattern were also sequenced at the UTK Sequencing Facility using primers 1055F and 1492R. In addition, the nearly fulllength rRNA gene of strain RD1 was sequenced with primers 27F, 530F (GTGCCAGCMGCCGCGG), 926F (AAACTYAAAKGAATTGACGG), and 907R (CCGTCAATTCMTTTRAGTTT) [24]. The partial sequences (each approx. 325 bp in length) were compared with existing sequences using BLASTX [25]. The nucleotide sequences for the six consortium strains were deposited in
Chloride in culture supernatants was determined turbidimetrically [17] by adding 40 Wl of 5 M phosphoric acid containing 0.1 M silver nitrate to 160 Wl of supernatant in a 96-well microtiter dish. The precipitate was allowed to settle for 10 min and was then read on a Microquant 96well scanning spectrophotometer (Bio-Tek Instruments, Winooski, VA, USA) at 595 nm. Absorbance readings were compared with those from standards which gave an r2 = 0.99 over the range of 0.1^2 mM. In the cases where absorbance of the supernatant exceeded the range of the standard curve, the supernatants were diluted with MSM and measured again. 2.6. DNA extraction and PCR ampli¢cation for genomic ¢ngerprinting Genomic DNA from the consortium and isolates was used for PCR ampli¢cation to generate genetic pro¢les. For the consortium, DNA was obtained from a crude lysate generated in a FP120 bead-beater (Bio 101, Carlsbad, CA, USA) set at 4.5 m s31 for 30 s. Genomic DNA was isolated from individual strains according to procedures outlined by Ausubel et al. [18]. BOX-PCR genomic ¢ngerprints for the consortium lysate and the isolated strains [19] were generated using the BOX A1R primer (CTACGGCAAGGCGACGCTGACG) under the following conditions: initial denaturation at 94³C for 3 min followed by 30 cycles of denaturation at 92³C for 45 s, annealing at 50³C for 60 s, and extension at 65³C for 6 min. An additional 6 min of extension at 65³C completed the program. The BOX-PCR products were separated in a 1.5% agarose gel after electrophoresis at 80 mV for 2 h. All PCR ampli¢cations were performed with a PTC200 DNA Engine thermal cycler (MJ Research, Watertown, MS, USA). Ribosomal intergenic spacer analysis (RISA) was applied to the consortium lysate and isolated strains using primers 1055F (ATGGCTGTCGTCAGCT) [20] and 23SR (GGGTTBCCCCATTCRG) [21]. Thermal cycler
2.7. Ampli¢cation and cloning of 16S rDNA
FEMSEC 1245 5-7-01
108
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
Fig. 1. Mineralization of [14 C]triclosan by the bacterial consortium growing on 500 mg l31 triclosan as sole source of carbon and energy (F), by strain Rd1 growing on YEPG (b), and killed consortium controls (E). Asterisk (*) represents uniformly 14 C-labeled ring.
GenBank under the accession numbers AF292237^ AF292242. 3. Results 3.1. Triclosan-degrading consortium and isolates After ¢ve serial transfers to MSMT, a bacterial consortium was obtained which was able to use triclosan as its sole source of carbon and energy. After approximately 13 days of growth on 500 mg l31 triclosan, almost 35% of the total 14 C-labeled triclosan was mineralized and captured as [14 C]CO2 (Fig. 1). Neither HPLC nor GC-MS analyses revealed the accumulation of any triclosan breakdown products in consortium supernatants. However, extraction of acidi¢ed culture supernatants with ethyl ether revealed that more than 95% of the added radiolabeled triclosan had disappeared (Table 1). A concomitant increase in the concentration of free chloride was also detected in supernatants after 13 days (Table 1). In broth culture, the consortium had a pinkish-red tint and reached stationary phase after approximately 4^6 days (Fig. 2). Repeated experiments demonstrated that mineralization proceeded without delay upon inoculation with 10% (v/v) of an exponential phase culture, however, active growth did not occur until after a lag of 1^2 days. When an inoculum volume of less than 5% (v/v) was transferred Table 1 Growing cultures and killed controls were analyzed after 13 days for total chloride release in the supernatant and for residual radioactivity in acidi¢ed ethyl ether extracts of the supernatant
Residual triclosan (%) Chloride released (%)
Growing cell culture
Killed control
4.5 (0.49) 87 (8.1)
98 (10) 10 (1.1)
Standard deviations are in parentheses.
Fig. 2. Growth of consortium (F) and killed controls (E) on 500 mg l31 triclosan as measured by increase in protein content.
to fresh £asks of MSMT broth, growth on triclosan was idiosyncratic, required long lag phases and was occasionally lost. No colonies were obtained when the consortium was plated onto either MSMT or MSMTYE agar. Five bacterial strains with morphologically distinct colony types were isolated after streaking the consortium on LB, YEPG, and PIA plates. None of these isolates, either separately or in combination, was able to utilize triclosan as a sole carbon source in liquid media. The isolates were unable to mineralize [14 C]triclosan when growing in pure or in de¢ned mixed cultures. Attempts were made to reduce the number of strains in the consortium by plating either dilute [15] or concentrated cell suspension onto MSMT, followed by resuspension of the plated organisms. After an extended incubation time of 3 weeks, the cells that were resuspended from the plates failed to restore consortium growth in MSMT broth [15]. 3.2. Assessment of uncultivated consortium members To address the possibility that additional strains were present in the triclosan-degrading consortium that were not isolated on the complex media plates, several molecular-genetic analyses were applied to the consortium lysate and to the ¢ve individual strains isolated from the consortium. The ampli¢ed DNA pro¢le generated by BOX-PCR demonstrated that none of the individual isolates had a genomic ¢ngerprint similar to that obtained from the consortium cell lysate (Fig. 3A). Although RISA pro¢les demonstrated the presence of similarly sized bands from both the isolates and the consortium, the predominance of a broad, poorly resolved 1400-bp band in the consortium ¢ngerprint suggested the presence of consortium members not represented by the ¢ve initially isolated strains (Fig. 3B). This hypothesis was supported by comparison of the ARDRA pro¢les from the consortium and the ¢ve isolate strains. Several bands from the consortium pro¢le were not associated with any of the isolates (Fig. 3C), suggest-
FEMSEC 1245 5-7-01
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
109
Fig. 3. Molecular analysis of isolates with distinct morphology from the triclosan-degrading consortium. A: BOX-PCR. B: RISA. C: ARDRA. Lane L is a 250-bp ladder from Gibco BRL in A and B, and the 100-bp ladder from Promega in C. Lane 1 represents strain Ly, 2 is strain Spd, 3 is strain Swo, 4 is strain Sws, 5 is strain Sy, and lane Cs is total consortium DNA. Arrows in C indicate bands unique to the consortium.
ing that additional consortium members were present and not accounted for by the ¢ve previously isolated strains. In order to gain a better understanding of which consortium members might be absent from the initial collection of strains isolated on complex media, 16S rRNA genes were ampli¢ed and cloned from the consortium lysate. Sequence analysis of 12 rDNA sequences randomly cloned from the consortium lysate revealed that four of the cloned 16S genes were identical, and came from an organism that had no signi¢cant identity to the partial 16S sequences from the ¢ve isolated strains. Sequence comparison done using the BLASTX program [25] showed this set of novel clones to be most similar to the Sphingomonas group of K-Proteobacteria. Of the eight remaining clones, ¢ve contained sequences identical to that of the strain Swo, while three contained sequences identical to strain Ly.
[14 C]CO2 and one strain, designated Rd1, was chosen for further study. Although strain Rd1 was able to mineralize 14 C-labeled triclosan when grown in complex medium containing 500 mg l31 triclosan (results not shown), HPLC and GC-MS analyses did not reveal the accumulation of any metabolites. Strain Rd1 was unable to grow on either MSMT/MSMTYE plates or broth. No growth was observed on minimal media containing acetate, succinate, glycerol, glucose, or benzoate as the sole carbon and energy source. When YEPG-grown Rd1 cells were washed, resuspended in MSM and incubated in a Biolog GN microtiter plate, none of the wells changed color during the ¢rst 2 weeks. After 2 weeks, a faint color change was observed in wells containing the amino acids arginine, proline and lysine. However, subsequent attempts at both
3.3. Isolation of a triclosan-degrading strain Previous experience with Sphingomonads suggested to us that slow growth might have been one of the reasons that no strains with similar 16S rRNA genes were isolated from the consortium. This was especially problematic given the spreading colony morphology of strain SPD, which tended to overgrow plates incubated for longer than 1 week at 28³C. To determine if some consortium members were being out-competed or overgrown during the initial isolation attempts, LB, PIA, and MSMA plates were streaked with an inoculum from the consortium and incubated at 22³C for 3 weeks. Incubation at this lower temperature dramatically reduced the spreading growth of strain SPD, and after 3 weeks a pair of small ( 6 1 mm) pink colonies were observed on PIA plates. After restreaking on PIA, these morphologically similar isolates were inoculated into YEPG broth amended with [14 C]triclosan and incubated at 28³C. Both isolates were able to liberate
Fig. 4. Comparison of (A) BOX-PCR, (B) RISA and (C) ARDRA patterns from strain Rd1 and the total consortium DNA. Lane L represent the same ladders used in Fig. 3. R represents strain Rd1 while Cs represent the consortium. Arrows in C represent bands unique to strain Rd1 and the consortium.
FEMSEC 1245 5-7-01
110
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
broth and plate cultivation suggested that strain Rd1 could not use any of these compounds as a sole source of carbon or energy for growth. 3.4. Phylogenetic analysis of consortium members BOX-PCR, RISA and ARDRA ¢ngerprinting of strain Rd1 showed bands that had been present in the consortium pro¢le but that had been missing from any of the other ¢ve isolate pro¢les (Fig. 4A^C). Sequencing of the 16S rRNA gene of strain Rd1 revealed that it was most closely related to a cluster of previously characterized KProteobacteria in the Sphingomonas group and was identical to the Sphingomonas-like partial sequences ampli¢ed and cloned from the consortium lysate. Sequence comparisons revealed that several of the partial 16S rDNA sequences from consortium isolates were 100% identical to the corresponding region of rDNA from previously cultivated organisms or rDNA sequences from uncultivated organism deposited in public databases. The partial rDNA sequences from strains Spd and Ly were each identical to a handful of sequences from bacteria, many of which were involved in the degradation of organic compounds including chlorinated solvents (Table 2). Strain Swo partial rDNA was identical to Alcaligenes sp. strain HR5 [26]. The remaining partial 16S sequences, from isolates Rd1, Sws and Sy, were not 100% identical to any sequence currently found in a public database. The identity of those partial sequences ranged from 98.5 to 99.1% (Table 2). 4. Discussion A stable enrichment culture was obtained that was able to use triclosan as a sole source of carbon and energy. This is the ¢rst report of a bacterial consortium able to use triclosan as a sole source of carbon and energy. Despite numerous attempts to isolate triclosan-degrading strains on solid MSMT, no growth was ever observed on those plates. Contrary to several recent reports [15,27,28], resuspension of cells from the initial dilution plates also failed to yield a single isolate that was capable of degrading triclosan in liquid medium. The initial failure to isolate pure strains from stable
enrichment cultures may indicate that cometabolic steps are required to degrade the substrate [29]. In the case of a recently characterized atrazine-degrading consortium [30], failure to isolate a single strain able to completely mineralize atrazine was due to distribution of the necessary catabolic genes between two di¡erent strains. In other instances, the organism responsible for the metabolic reaction of interest may be fastidious, requiring cofactors from other consortium members [31]. Our inability to cultivate strain Rd1 on minimal media containing either triclosan or other single carbon sources suggests that this strain may fall into this last category, requiring cofactors supplied only in the consortium or complex media. The apparent lag in growth despite immediate mineralization of triclosan (Figs. 1 and 2) suggests that triclosan degradation may be a detoxi¢cation mechanism. This hypothesis is supported by the isolation of Rd1M, a mutant of Rd1 that is impaired in its ability to mineralize triclosan [32]. While Rd1 readily grows on YEPG containing 500 mg l31 triclosan, the growth of strain Rd1M is inhibited by triclosan concentrations in excess of 50 mg l31 . Further work needs to be done to determine what e¡ect degradation has on the overall resistance of Rd1 to triclosan. Despite clear evidence for the partial mineralization of the 14 C-labeled dichloro ring by the consortium and by strain Rd1, no triclosan metabolites were detected by either GC-MS or HPLC analysis of culture supernatants. Previous work by others on the bacterial degradation of various substituted and unsubstituted diphenyl ethers suggests that degradation may occur by either of two mechanisms. One mechanism involves dihydroxylation of one of the rings in the 2,3 positions followed by either intradiol [33] or extradiol [34,35] cleavage. The second mechanism requires a highly regioselective, angular dioxygenation at the 1,2 positions of one of the rings [36], which results in the spontaneous ¢ssion of the ether bond, yielding both a phenol and a catechol moiety [37^40]. Further work is needed to determine whether strain Rd1 relies on either of these, or another, as yet unde¢ned mechanism for the degradation of triclosan. A recent report by Hundt et al. identi¢ed dichlorophenol and several glycosylated triclosan derivatives as transformation products of triclosan by the fungal species Pycnoporus cinnabarinus and Trametes versicolor [41]. Although we were able to detect 2,4-dichlorophenol in
Table 2 Accession numbers and closest 16S match for organisms isolated from the triclosan-degrading consortium Consortium isolate (accession number)
Closest 16S match (accession number)
Ly Spd Swo Sy Sws Rd1
Pseudomonas mendocina Pseudomonas aeruginosa Alcaligenes xylosoxidans Rhodanobacter lindanoclasticus Agrobacterium radiobacter Sphingomonas sanguis
(AF292237) (AF292239) (AF292240) (AF292242) (AF292241) (AF292238)
FEMSEC 1245 5-7-01
Identity (%) (AF232713) (AB037560) (AJ002808) (AF039167) (D137260) (AF226204)
100 100 100 99.1 98.8 98.5
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
analytical standards down to 100 ng l31 via HPLC, there was no evidence for 2,4-dichlorophenol production either from the consortium or from strain Rd1 individually. When strain Rd1 was grown on complex media, the extent of 14 C-labeled triclosan mineralization was similar to that of the consortium growing on minimal media (data not shown). This suggests that Rd1 is responsible for the bulk of the triclosan mineralization observed and that none of the other consortium members can signi¢cantly degrade the remaining labeled triclosan residues. However, this does not preclude the participation of other consortium members in the degradation of unlabeled, monochloro ring, as total chloride release of greater than 85% indicates signi¢cant transformation of both rings (Table 1). While the partial 16S rDNA sequences obtained from the six isolates are insu¤cient for a detailed phylogenetic determination, it is nevertheless clear that each strain can be assigned to a speci¢c subgroup among the K (strains Rd1 and Sws), L (strain Swo) or Q (strains Spd, Ly, and Sy) Proteobacterial subdivisions. Particularly noteworthy among the isolates is strain Sy, which represents one of only two cultivated members of the Rhodanobacter lindanoclasticus subgroup of the Q-Proteobacteria [42]. R. lindanoclasticus is a novel species that was isolated using the insecticide and wood preservative lindane (Q-hexachlorocyclohexane) as a sole carbon source. Although strain Sy appears to be phylogenetically related to organisms involved in the degradation of chlorinated organic compounds, its role and the role of the other consortium members in the mineralization of triclosan is unclear. Strains from several di¡erent genera have been described which are able to degrade various diphenyl ethers. These genera include Erwinia [43], Pseudomonas [33,35,44], and Sphingomonas [38]. It is interesting to note that only Sphingomonas-like strains have thus far been reported to degrade chlorinated diphenyl ethers [38,39]. Strain Rd1 is the ¢rst organism reported to partially mineralize triclosan. The isolation of strain Rd1 was guided in large part by the sequence information obtained through random cloning and sequencing of partial 16S rRNA gene fragments from the consortium DNA extract. Teske et al. [45] used 16S rDNA sequence information to design selective enrichment conditions for the isolation of individual organisms from a mixed culture. Other molecular-genetic techniques such as ARDRA and REP-PCR have also been used to guide the isolation of refractory organisms from mixed cultures [15]. This work adds to the evidence demonstrating the value of molecular ecology techniques in the characterization of microbial consortia [15,45]. Acknowledgements This work was supported in part by a Dow Foundation SPHERE award to G.S.S. and in part by the Waste Man-
111
agement Research and Education Institute, University of Tennessee, Knoxville. A.G.H. was supported by an appointment to the Alexander Hollaender Distinguished Postdoctoral Fellowship Program sponsored by the US Department of Energy, O¤ce of Health and Environmental Research, and administered by the Oak Ridge Institute for Science and Education. The authors wish to thank Josef Inauen of Ciba Specialty Chemical for the generous gifts of labeled and unlabeled triclosan.
References [1] McMurry, L.M., Oethinger, M. and Levy, S.B. (1998) Triclosan targets lipid synthesis. Nature 394, 531^532. [2] Daughton, C.G. and Ternes, T.A. (1999) Pharmaceuticals and personal care products in the environment : agents of subtle change? Environ. Health Perspect. 107, 907^937. [3] Parikh, S., Xiao, G. and Tonge, P.J. (2000) Inhibition of InhA, the enoyl reductase from mycobacterium tuberculosis, by triclosan and isoniazid. Biochemistry 39, 7645^7650. [4] Heath, R.J. and Rock, C.O. (2000) A triclosan-resistant bacterial enzyme. Nature 406, 145^146. [5] Schweizer, H.P. (1998) Intrinsic resistance to inhibitors of fatty acid biosynthesis in Pseudomonas aeruginosa is due to e¥ux: application of a novel technique for generation of unmarked chromosomal mutations for the study of e¥ux systems. Antimicrob. Agents Chemother. 42, 394^398. [6] Difco Laboratories (1985) Difco Laboratories Incorporated, Detroit, MI. [7] Voets, J.P., Pipyn, P., Van Lancker, P. and Verstraete, W. (1976) Degradation of microbicides under di¡erent environmental conditions. J. Bacteriol. 40, 67^72. [8] Lopez-Avila, V. and Hites, R.A. (1980) Organic compounds in an industrial wastewater : their transport into sediments. Environ. Sci. Technol. 14, 1382^1390. [9] Hale, R.C., Smith, C.L., De Fur, P.O., Harvey, E., Bush, E.O., La Guardia, M.J. and Vadas, G.G. (2000) Nonylphenols in sediments and e¥uents associated with diverse wastewater outfalls. Environ. Toxicol. Chem. 19, 946^952. [10] Okumura, T. and Nishikawa, Y. (1996) Gas chromatography-mass spectrometry determination of trilcosans in water, sediment, and ¢sh samples via methylation with diazomethane. Anal. Chim. Acta 325, 175^184. [11] Miyazaki, T., Yamagishi, T. and Matsumoto, M. (1984) Residues of 4-chloro-1-(2,4-dichlorophenoxy)-2-methoxybenzene (triclosan methyl) in aquatic biota. Bull. Environ. Contam. 32, 227^232. [12] Beck, H.A.D., Eckart, K., Mathar, W. and Wittkowski, R. (1989) Determination of PCDDs and PCDFs in Irgasan DP 300. Chemosphere 19, 167^170. [13] McCullar, M.V., Brenner, V.H.A.R. and Focht, D.D. (1994) Construction of a novel polychlorinated biphenyl-degrading bacterium: Utilization of 3,4P-dichlorobiphenyl by Pseudomonas acidovorans M3GY. Appl. Environ. Microbiol. 60, 3833^3839. [14] Arensdorf, J.J. and Focht, D.D. (1995) A meta cleavage pathway for 4-chlorobenzoate, an intermediate in the metabolism of 4-chlorobiphenyl by Pseudomonas cepacia P166. Appl. Environ. Microbiol. 61, 443^447. [15] Maltseva, O. and Oriel, P. (1997) Monitoring of an alkaline 2,4,6trichlorophenol-degrading enrichment culture by DNA ¢ngerprinting methods and isolation of the responsible organism, haloalkiphilic Nocardioides sp. strain M6. Appl. Environ. Microbiol. 63, 4145^ 4149. [16] Hay, A.G. and Focht, D.D. (1998) Cometabolism of 1,1-dichloro-
FEMSEC 1245 5-7-01
112
[17]
[18]
[19]
[20]
[21]
[22] [23]
[24]
[25]
[26]
[27]
[28]
[29]
[30]
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112 2,2-bis(4-chlorophenyl)ethylene by Pseudomonas acidovorans M3GY grown on biphenyl. Appl. Environ. Microbiol. 64, 2141^2146. Focht, D.D. and Alexander, M. (1971) Aerobic cometabolism of DDT analogues by Hydrogenomonas sp.. J. Agric. Food Chem. 19, 20^22. Ausubel, F.M., Brent, R., Kingston, R.E., More, D.D., Seidman, J.G., Smith, J.A. and Struhl, K. (1989) Current protocols in Molecular Biology John Wiley and Sons, New York, NY. Rademaker, J.L.W., Louws, F.J. and De Bruijn, F.J. (1998) Characterization of the diversity of ecologically important microbes by repPRC genomic ¢ngerprinting in: Molecular Microbiology Ecology Manual, Vol. 3 (Akkermans, A.D.L., Van Elsas, J.D. and De Bruijn, F.J., Eds.). Kluwer Academic, Dordrecht. Ferris, M.J., Muyzer, G. and Ward, D.M. (1996) Denaturing gradient gel electrophoresis pro¢les of 16S rRNA-de¢ned populations inhabiting a hot spring microbial mat community. Appl. Environ. Microbiol. 62, 340^346. Borneman, J. and Triplett, E.W. (1997) Molecular microbial diversity in soils from eastern Amazonia: Evidence for unusual microorganisms and microbial population shifts associated with deforestation. Appl. Environ. Microbiol. 63, 2647^2653. Roux, K.H. (1994) Using mismatched primer-template pairs in touchdown PCR. BioTechniques 16, 812^814. Vaneechoutte, M. et al. (1992) Rapid identi¢cation of bacteria of the Comamonadaceae with ampli¢ed ribosomal DNA-restriction analysis (ARDRA). FEMS Microbiol. Lett. 93, 227^234. Lane, D.J. (1991) 16S/235 rRNA sequencing in: Nucleic Acid Techniques in Bacterial Systematics (Stackebrandt, E. and Goodfellow, M., Eds.), pp. 115^147. John Wiley and Sons, New York. Altschul, S.F., Madden, T.L., Scha¡er, A.A., Zhang, J., Zhang, Z., Miller, W. and Lipman, D.J. (1997) Gapped BLAST and PSIBLAST: a new generation of protein database search programs. Nucleic Acids Res. 25, 3389^3402. Ho¡mann, A., Thimm, T., Droege, M., Moore, E.R.B., Munch, J.C. and Tebbe, C.C. (1998) Intergeneric transfer of conjugative and mobilizable plasmids harbored by Escherichia coli in the gut of the soil microarthropod Folsomia candida (Collembola). Appl. Environ. Microbiol. 64, 2652^2659. Sutherland, T.D., Horne, I., Lacey, M.J., Harcourt, R.L., Russel, R.J. and Oakeshott, J.G. (2000) Enrichment of an endosulfane-degrading mixed bacterial culture. Appl. Environ. Microbiol. 66, 2822^ 2828. Tanghe, T., Dhooge, W. and Verstraete, W. (1999) Isolation of a bacterial strain able to degrade branched nonylphenol. Appl. Environ. Microbiol. 65, 746^751. Focht, D.D. (1993) in: Soil Biochemistry, Vol. 8 (Bollag, M. and Stozky, G., Eds.). Marcel Dekker, New York Microbial degradation of chlorimated biphenyls, 341^407. De Souza, M.L., Newcombe, D., Alvey, S., Crowley, D., Hay, A., Sadowsky, M.J. and Wackett, L.P. (1998) Molecular basis of a bac-
[31]
[32] [33]
[34]
[35]
[36]
[37]
[38]
[39]
[40]
[41]
[42] [43]
[44]
[45]
terial consortium: interspecies catabolism of atrazine. Appl. Environ. Microbiol. 64, 178^184. Maymo-Gatell, X., Chien, Y.-T., Gossett, J.M. and Zinder, S.H. (1997) Isolation of a bacterium that reductively dechlorinates tetrachloroethylene to ethene. Science 276, 1568^1571. Hay, A.G., Unpublished results. Takase, I., Omori, T. and Minoda, Y. (1986) Microbial degradation products from biphenyl-related compounds. Agric. Biol. Chem. 50, 681^686. Pfeifer, F., Trueper, H.-G., Klein, J. and Schacht, S. (1993) Degradation of diphenylether by Pseudomonas cepacia Et4: Enzymatic release of phenol from 2,3-dihydroxydiphenyl ether. Arch. Microbiol. 159, 323^329. Pfeifer, F., Schacht, S., Klein, J. and Truper, H.G. (1989) Degradation of diphenylether by Pseudomonas cepacia. Arch. Microbiol. 152, 515^519. Fortnagel, P., Wittich, R.M., Harms, H., Schmidt, S., Franke, S., Sinnwell, V., Wilkes, H. and Francke, W. (1989) New bacterial degradation of the biaryl ether structure regioselective dioxygenation prompts cleavage of ether bonds. Naturwissenschaften 76, 523^524. Schmidt, S., Wittich, R.M., Fortnagel, P., Erdmann, D. and Francke, W. (1992) Metabolism of 3-methyldiphenyl ether by Sphingomonas sp. SS31. FEMS Microbiol. Lett. 96, 253^258. Schmidt, S., Wittich, R.M., Erdmann, D., Wilkes, H., Francke, W. and Fortnagel, P. (1992) Biodegradation of diphenyl ether and its monohalogenated derivatives by Sphingomonas sp. strain SS3. Appl. Environ. Microbiol. 58, 2744^2750. Schmidt, S., Fortnagel, P. and Wittich, R.M. (1993) Biodegradation and transformation of 4,4P- and 2,4-dihalodiphenyl ethers by Sphingomonas sp. strain SS33. Appl. Environ. Microbiol. 59, 3931^3933. Wilkes, H., Francke, W., Wittich, R.M., Harms, H., Schmidt, S. and Fortnagel, P. (1992) Mechanistic investigations on microbial degradation of diaryl ethers analysis of isotope-labeled reaction products. Naturwissenschaften 79, 269^271. Hundt, K., Martin, D., Hammer, E., Jonas, U., Kindermann, M.K. and Schauer, F. (2000) Transformation of triclosan by Trametes versicolor and Pycnoporus cinnabarinus. Appl. Environ. Microbiol. 66, 4157^4160. Maidak, B.L. et al. (2000) The RDP (Ribosomal Database Project) continues. Nucleic Acids Res. 28, 173^174. Liaw, J.H. and Srinivasan (1989) Molecular cloning and expression of an Erwinia sp. gene encoding diphenyl ether cleavage in Escherichia coli. Appl. Environ. Microbiol. 55, 2220^2225. Wittich, R.M., Schmidt, S. and Fortnagel, P. (1990) Bacterial degradation of 3- and 4-carboxybiphenyl ether by Pseudomonas sp. NSS2. FEMS Microbiol. Lett. 67, 157^160. Teske, A.P.S., Cohen, Y. and Muyzer, G. (1996) Molecular identi¢cation of bacteria from a coculture by denaturing gradient gel electrophorsis of 16S ribosomal DNA fragments as a tool for isolation in pure cultures. Appl. Environ. Microbiol. 62, 4210^4215.
FEMSEC 1245 5-7-01
www.fems-microbiology.org
Growth of a bacterial consortium on triclosan Anthony G. Hay a
a;b;c
, Peter M. Dees b , Gary S. Sayler
a;
*
Center for Environmental Biotechnology, 676 Dabney Hall, University of Tennessee, Knoxville, TN 37996-1605, USA b Department of Microbiology, Cornell University, Ithaca, NY 14853, USA c Institute for Comparative and Environmental Toxicology, Cornell University, Ithaca, NY 14853, USA Received 26 September 2000; received in revised form 6 April 2001; accepted 8 April 2001 First published online 21 May 2001
Abstract Triclosan is a polychlorinated hydroxy diphenylether that has been widely used as an antimicrobial compound. An enrichment using triclosan as a sole source of carbon and energy yielded a consortium of bacteria capable of growing on this compound. The dichloro ring was partially mineralized, resulting in the conversion of approximately 35% of the [14 C]triclosan to [14 C]CO2 . Use of molecular fingerprinting techniques and 16S rDNA cloning and sequencing aided in the identification and eventual isolation of an auxotrophic Sphingomonas-like organism, strain Rd1, which was able to partially mineralize triclosan when grown on complex media. ß 2001 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. Keywords : Biodegradation; Consortium ; Sphingomonas ; Triclosan; Antimicrobial
1. Introduction Triclosan is a broad spectrum antimicrobial compound that has recently found increased use in personal care products such as soaps, handcreams, toothpastes, and acne creams and in commercial products such as hosiery and plastics [1,2]. Evidence suggests that triclosan blocks fatty acid synthesis by interacting with the NADP binding site of the enoyl-ACP reductase, FabI in Escherichia coli [1] and InhA in Mycobacterium tuberculosis [3]. However, Streptococcus pneumoniae, which is also sensitive to triclosan, does not contain a FabI homolog [4]. Instead S. pneumoniae contains FabK, an isofunctional FAD-dependent enoyl-ACP reductase which is not sensitive to triclosan [4]. This suggests that triclosan must have another target in S. pneumoniae [4]. Pseudomonas aeruginosa, which is highly resistant to triclosan, contains both FabI and FabK, in addition to a multidrug e¥ux pump which is able to transport triclosan out of the cell [5]. Triclosan, otherwise known as Irgasan, is the selective ingredient in Pseudomonas Isolation Agar (PIA) [6].
* Corresponding author. Tel. : +1 (423) 974-8080; Fax: +1 (423) 974-8086; E-mail: [email protected]
Early work done by Voets et al. [7] demonstrated that up to 50% of the triclosan added to synthetic sewage sludge was degraded under both aerobic and anaerobic conditions via cometabolism. In minimal media, however, no degradation was observed. The authors therefore suggested that triclosan might persist in oligotrophic waters if it had not ¢rst been subjected to e¤cient wastewater treatment procedures [7]. Although there is little evidence to suggest widespread environmental accumulation or recalcitrance, several studies have demonstrated the presence of triclosan in sediments [8,9] and biota [10,11]. Concern about the environmental toxicity of the compound comes in part from the predioxin nature of triclosan which, upon exposure to heat or radiation, can undergo ring closure to form chlorinated dioxins [10]. This concern is compounded by the fact that di- and trichlorinated dibenzodioxins have been shown to contaminate commercial triclosan preparations [12]. Given concern about the widespread use of antimicrobial compounds and the possible emergence of drug-resistant bacteria, we sought to assess the potential for biodegradation of triclosan by bacteria in activated sludge from a residential wastewater treatment facility. In this study we describe the isolation and phylogenetic characterization of six strains from a consortium of bacteria able to grow on triclosan as a sole source of carbon and energy.
0168-6496 / 01 / $20.00 ß 2001 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. PII: S 0 1 6 8 - 6 4 9 6 ( 0 1 ) 0 0 1 2 7 - 1
FEMSEC 1245 5-7-01
106
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
2. Materials and methods 2.1. Bacteriological media Enrichments and subsequent cultivation of the consortium were done in a de¢ned, chloride-free minimal salts medium (MSM) containing 10 mM K2 HPO4 , 3 mM NaHPO4 , 10 mM (NH4 )2 SO4 , and 1 mM MgSO4 in deionized water. A chloride-free trace elements solution was also added at 1 ml l31 [13]. Consortium members were also streaked on MSM plates containing 1000 mg l31 sodium acetate (MSMA), 500 mg l31 triclosan as a sole source of carbon and energy (MSMT), with and without 50 mg l31 yeast extract (MSMTYE) and on the following complex media: Luria^Bertani medium (LB), PIA (Difco, Detroit, MI, USA) and YEPG. YEPG consists of 0.2 g of yeast extract, 2 g of peptone, 0.2 g of ammonium nitrate, and 1 g of glucose in 1 l of distilled water [14]. All solid media were solidi¢ed with 15 g l31 bacteriological agar or noble agar in the case of the minimal media. 2.2. Reagents Triclosan (5-chloro-2-(2,4-dichlorophenoxy)phenol) and [ C]triclosan were a gift from Ciba Specialty Chemical (Basel, Switzerland) and were 99% and s 99% pure respectively. The [14 C]triclosan had a speci¢c activity of 5.2 MBq mg31 and was uniformly labeled in the dichloro ring, while the other ring was unlabeled. Reagent grade ethyl ether, acetonitrile and Scintisafe Econo 1 scintillation cocktail were obtained from Fisher Scienti¢c (Pittsburgh, PA, USA). N-Nitrosyl-N-methylurea, which was used to generate diazomethane [14], was obtained from Sigma Chemical Company (St. Louis, MO, USA). 14
2.3. Bacterial cultures and conditions Activated sludge from the Neyland Wastewater treatment plant (Knoxville, TN, USA) was used as a source of inoculum for enrichments on triclosan. Initially, 500 mg of powdered triclosan was added directly to 1 l of activated sludge and incubated for 1 month at 28³C while being shaken at 150 rpm. After the initial 1 month incubation and subsequently every 2 weeks thereafter, the enrichment culture was transferred (10% v/v) to fresh MSM containing 500 mg l31 triclosan. Two weeks after the ¢fth transfer, the enrichment was streaked onto all of the media listed above and plates were incubated at 28³C. Individual colonies with distinctive morphologies were picked from the complex media plates and streaked on the same medium for single colony isolation, then restreaked on MSMT and MSMTYE plates. The enrichment culture was serially diluted and plated on MSMT and MSMTYE agar plates, which were then incubated for 1^3 weeks at room temperature. After incubation, the plates were wiped with a presoaked sterile cot-
ton swab, resuspended in 100 ml MSMT, incubated at 28³C and examined for an increase in visible turbidity [15]. In addition, to assess the possibility that numerically inferior organisms were required for triclosan mineralization, aliquots amounting to 10 ml of enrichment culture were centrifuged at 8000Ug in a microcentrifuge, the supernatant was decanted and the combined cell pellets were resuspended in 0.1 ml 30 mM phosphate bu¡er (pH 7.3). The 100U concentrated cell suspension was spread on MSMT and MSMTYE plates and incubated at 28³C for as long as 1 month. The plates were then wiped as above and resuspended in MSMT to check for growth. When grown in MSMT broth, the consortium often clumped. To avoid variation when optical density was used as a measure of growth, the growth of the consortium was monitored as an increase in protein content. The enrichment culture was vigorously shaken before sampling to disperse clumped cells and after dispersal 250-Wl aliquots of culture were transferred to individual 1.5-ml microcentrifuge tubes and frozen until the end of the experiment. Frozen cells were thawed, then permeabilized by adding 5 Wl of toluene and the protein concentration was measured using the Bio-Rad Protein Assay kit (Bio-Rad, Richmond, CA, USA) according to the manufacturer's instructions. 2.4. Mineralization assay and metabolite analysis The consortium and individual isolates were screened for triclosan mineralization by placing 5 ml of a triclosan-grown (consortium) or YEPG-grown, exponential phase (isolates) culture in a 40-ml EPA vial containing a smaller 8-ml glass vial to which 1 ml of 1 M NaOH was added. A mixture of unlabeled and 14 C-labeled triclosan in acetone was added to the culture to give a ¢nal concentration of 500 mg l31 and 250 000 dpm. The 40-ml EPA vials were then capped and shaken at 150 rpm while incubating at 28³C. The full volume of spent NaOH was periodically withdrawn and replaced with a fresh 1.0 ml of 1 M NaOH. The spent NaOH was added to 10 ml scintillation cocktail and [14 C]CO2 accumulation was measured using a Beckman LS5000CE liquid scintillation counter (Beckman, Fullerton, CA, USA). No growth was observed when acetone was added as a sole source of carbon and energy. After the cells were pelleted by microcentrifugation, the consortium supernatant was analyzed for the accumulation of metabolites by high pressure liquid chromatography (HPLC) using a Supelcosil LC-18-T reverse phase C18 column (Supelco, Bellefonte, PA, USA) attached to a binary LC 250 series pump with a model LC-235 diode array detector (both Perkin Elmer, Foster City, CA, USA). Absorbance was monitored at 210 nm. The eluent was 50% 40 mM acetic acid and 50% acetonitrile. Samples of the culture supernatant were also analyzed by gas chro-
FEMSEC 1245 5-7-01
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
107
matography. After centrifuging 100 ml of culture (10 000Ug for 10 min), the supernatant was acidi¢ed with HCl and then extracted three times with ethyl ether. The extracts were pooled, evaporated to dryness, then resuspended in 1.0 ml of ethyl ether and either derivatized with diazomethane [16] or analyzed in the underivatized state. Analysis of extracts was performed on a Hewlett Packard 6890 series gas chromatograph equipped with an HP5ms column (30 mU250 WmU0.25 Wm) and a 5973 series mass selective detector (GC-MS) (Hewlett Packard, Wilmington, DE, USA). The injector was held at 250³C. The initial oven temperature was 40³C and was ramped after 1 min at 10³C min31 to 250³C at which point the temperature was held constant for 10 min.
conditions were based on a modi¢ed touchdown protocol [22]. The initial denaturation step for the ¢rst primer set was 5 min at 95³C followed by 10 cycles of denaturation at 95³C for 45 s, annealing at 65³C for 15 s, and extension at 70³C for 60 s. For touchdown PCR, the annealing temperature was decreased by 1³C cycle31 for the ¢rst 10 cycles, followed by 20 cycles of annealing at 55³C. A 3-min extension at 70³C was included after completion of the 30 cycles of touchdown ampli¢cation. PCR products ampli¢ed from the ribosomal intergenic spacer region were separated in a 2% agarose gel after electrophoresis for 4 h at 80 mV.
2.5. Chloride determination
PCR was also used to amplify nearly full-length sections of the 16S rRNA gene from the consortium DNA lysate and individual isolates using universal primers 27F (AGAGTTTGATCMTGGCTCAG) and 1492R (TACGGYTACCTTGTTACGACTT). Touchdown thermal cycling conditions for the 27F/1492R primer set were similar to the protocol for RISA PCR except that the initial annealing temperatures for the ¢rst 10 cycles decreased at 1³C cycle31 from 55³C to 45³C, followed by 20 cycles of annealing at 45³C. PCR products ampli¢ed with the 27F/1492R primer pair were ligated into pCR2.1-TOPO (Invitrogen, Carlsbad, CA, USA) following the manufacturer's instructions. The ligation mixture was electroporated into DH5K electrocompetent cells and plated onto LB containing 50 mg l31 kanamycin and 40 mg ml31 X-gal. Plasmids were puri¢ed from white transformant colonies using Mini Prep Express Matrix (Bio 101, Carlsbad, CA, USA) following the manufacturer's instructions and plasmid DNA was screened for the presence of a 1500-bp insert after digestion with EcoRI and electrophoresis of the plasmid DNA in a 1% agarose gel. After screening for inserts, plasmids from 12 of the transformants were used for sequencing templates at the University of Tennessee, Knoxville (UTK) Sequencing Facility using primer 1492R. PCR products ampli¢ed directly from the consortium DNA extract and from individual isolates using the 27F/ 1492R primer set were digested with HhaI and separated in a 4% agarose gel after electrophoresis for 12 h at 25 mV. The rRNA genes from ¢ve isolates which gave either a unique BOX, RISA, or ampli¢ed ribosomal DNA restriction analysis (ARDRA) [23]) pattern were also sequenced at the UTK Sequencing Facility using primers 1055F and 1492R. In addition, the nearly fulllength rRNA gene of strain RD1 was sequenced with primers 27F, 530F (GTGCCAGCMGCCGCGG), 926F (AAACTYAAAKGAATTGACGG), and 907R (CCGTCAATTCMTTTRAGTTT) [24]. The partial sequences (each approx. 325 bp in length) were compared with existing sequences using BLASTX [25]. The nucleotide sequences for the six consortium strains were deposited in
Chloride in culture supernatants was determined turbidimetrically [17] by adding 40 Wl of 5 M phosphoric acid containing 0.1 M silver nitrate to 160 Wl of supernatant in a 96-well microtiter dish. The precipitate was allowed to settle for 10 min and was then read on a Microquant 96well scanning spectrophotometer (Bio-Tek Instruments, Winooski, VA, USA) at 595 nm. Absorbance readings were compared with those from standards which gave an r2 = 0.99 over the range of 0.1^2 mM. In the cases where absorbance of the supernatant exceeded the range of the standard curve, the supernatants were diluted with MSM and measured again. 2.6. DNA extraction and PCR ampli¢cation for genomic ¢ngerprinting Genomic DNA from the consortium and isolates was used for PCR ampli¢cation to generate genetic pro¢les. For the consortium, DNA was obtained from a crude lysate generated in a FP120 bead-beater (Bio 101, Carlsbad, CA, USA) set at 4.5 m s31 for 30 s. Genomic DNA was isolated from individual strains according to procedures outlined by Ausubel et al. [18]. BOX-PCR genomic ¢ngerprints for the consortium lysate and the isolated strains [19] were generated using the BOX A1R primer (CTACGGCAAGGCGACGCTGACG) under the following conditions: initial denaturation at 94³C for 3 min followed by 30 cycles of denaturation at 92³C for 45 s, annealing at 50³C for 60 s, and extension at 65³C for 6 min. An additional 6 min of extension at 65³C completed the program. The BOX-PCR products were separated in a 1.5% agarose gel after electrophoresis at 80 mV for 2 h. All PCR ampli¢cations were performed with a PTC200 DNA Engine thermal cycler (MJ Research, Watertown, MS, USA). Ribosomal intergenic spacer analysis (RISA) was applied to the consortium lysate and isolated strains using primers 1055F (ATGGCTGTCGTCAGCT) [20] and 23SR (GGGTTBCCCCATTCRG) [21]. Thermal cycler
2.7. Ampli¢cation and cloning of 16S rDNA
FEMSEC 1245 5-7-01
108
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
Fig. 1. Mineralization of [14 C]triclosan by the bacterial consortium growing on 500 mg l31 triclosan as sole source of carbon and energy (F), by strain Rd1 growing on YEPG (b), and killed consortium controls (E). Asterisk (*) represents uniformly 14 C-labeled ring.
GenBank under the accession numbers AF292237^ AF292242. 3. Results 3.1. Triclosan-degrading consortium and isolates After ¢ve serial transfers to MSMT, a bacterial consortium was obtained which was able to use triclosan as its sole source of carbon and energy. After approximately 13 days of growth on 500 mg l31 triclosan, almost 35% of the total 14 C-labeled triclosan was mineralized and captured as [14 C]CO2 (Fig. 1). Neither HPLC nor GC-MS analyses revealed the accumulation of any triclosan breakdown products in consortium supernatants. However, extraction of acidi¢ed culture supernatants with ethyl ether revealed that more than 95% of the added radiolabeled triclosan had disappeared (Table 1). A concomitant increase in the concentration of free chloride was also detected in supernatants after 13 days (Table 1). In broth culture, the consortium had a pinkish-red tint and reached stationary phase after approximately 4^6 days (Fig. 2). Repeated experiments demonstrated that mineralization proceeded without delay upon inoculation with 10% (v/v) of an exponential phase culture, however, active growth did not occur until after a lag of 1^2 days. When an inoculum volume of less than 5% (v/v) was transferred Table 1 Growing cultures and killed controls were analyzed after 13 days for total chloride release in the supernatant and for residual radioactivity in acidi¢ed ethyl ether extracts of the supernatant
Residual triclosan (%) Chloride released (%)
Growing cell culture
Killed control
4.5 (0.49) 87 (8.1)
98 (10) 10 (1.1)
Standard deviations are in parentheses.
Fig. 2. Growth of consortium (F) and killed controls (E) on 500 mg l31 triclosan as measured by increase in protein content.
to fresh £asks of MSMT broth, growth on triclosan was idiosyncratic, required long lag phases and was occasionally lost. No colonies were obtained when the consortium was plated onto either MSMT or MSMTYE agar. Five bacterial strains with morphologically distinct colony types were isolated after streaking the consortium on LB, YEPG, and PIA plates. None of these isolates, either separately or in combination, was able to utilize triclosan as a sole carbon source in liquid media. The isolates were unable to mineralize [14 C]triclosan when growing in pure or in de¢ned mixed cultures. Attempts were made to reduce the number of strains in the consortium by plating either dilute [15] or concentrated cell suspension onto MSMT, followed by resuspension of the plated organisms. After an extended incubation time of 3 weeks, the cells that were resuspended from the plates failed to restore consortium growth in MSMT broth [15]. 3.2. Assessment of uncultivated consortium members To address the possibility that additional strains were present in the triclosan-degrading consortium that were not isolated on the complex media plates, several molecular-genetic analyses were applied to the consortium lysate and to the ¢ve individual strains isolated from the consortium. The ampli¢ed DNA pro¢le generated by BOX-PCR demonstrated that none of the individual isolates had a genomic ¢ngerprint similar to that obtained from the consortium cell lysate (Fig. 3A). Although RISA pro¢les demonstrated the presence of similarly sized bands from both the isolates and the consortium, the predominance of a broad, poorly resolved 1400-bp band in the consortium ¢ngerprint suggested the presence of consortium members not represented by the ¢ve initially isolated strains (Fig. 3B). This hypothesis was supported by comparison of the ARDRA pro¢les from the consortium and the ¢ve isolate strains. Several bands from the consortium pro¢le were not associated with any of the isolates (Fig. 3C), suggest-
FEMSEC 1245 5-7-01
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
109
Fig. 3. Molecular analysis of isolates with distinct morphology from the triclosan-degrading consortium. A: BOX-PCR. B: RISA. C: ARDRA. Lane L is a 250-bp ladder from Gibco BRL in A and B, and the 100-bp ladder from Promega in C. Lane 1 represents strain Ly, 2 is strain Spd, 3 is strain Swo, 4 is strain Sws, 5 is strain Sy, and lane Cs is total consortium DNA. Arrows in C indicate bands unique to the consortium.
ing that additional consortium members were present and not accounted for by the ¢ve previously isolated strains. In order to gain a better understanding of which consortium members might be absent from the initial collection of strains isolated on complex media, 16S rRNA genes were ampli¢ed and cloned from the consortium lysate. Sequence analysis of 12 rDNA sequences randomly cloned from the consortium lysate revealed that four of the cloned 16S genes were identical, and came from an organism that had no signi¢cant identity to the partial 16S sequences from the ¢ve isolated strains. Sequence comparison done using the BLASTX program [25] showed this set of novel clones to be most similar to the Sphingomonas group of K-Proteobacteria. Of the eight remaining clones, ¢ve contained sequences identical to that of the strain Swo, while three contained sequences identical to strain Ly.
[14 C]CO2 and one strain, designated Rd1, was chosen for further study. Although strain Rd1 was able to mineralize 14 C-labeled triclosan when grown in complex medium containing 500 mg l31 triclosan (results not shown), HPLC and GC-MS analyses did not reveal the accumulation of any metabolites. Strain Rd1 was unable to grow on either MSMT/MSMTYE plates or broth. No growth was observed on minimal media containing acetate, succinate, glycerol, glucose, or benzoate as the sole carbon and energy source. When YEPG-grown Rd1 cells were washed, resuspended in MSM and incubated in a Biolog GN microtiter plate, none of the wells changed color during the ¢rst 2 weeks. After 2 weeks, a faint color change was observed in wells containing the amino acids arginine, proline and lysine. However, subsequent attempts at both
3.3. Isolation of a triclosan-degrading strain Previous experience with Sphingomonads suggested to us that slow growth might have been one of the reasons that no strains with similar 16S rRNA genes were isolated from the consortium. This was especially problematic given the spreading colony morphology of strain SPD, which tended to overgrow plates incubated for longer than 1 week at 28³C. To determine if some consortium members were being out-competed or overgrown during the initial isolation attempts, LB, PIA, and MSMA plates were streaked with an inoculum from the consortium and incubated at 22³C for 3 weeks. Incubation at this lower temperature dramatically reduced the spreading growth of strain SPD, and after 3 weeks a pair of small ( 6 1 mm) pink colonies were observed on PIA plates. After restreaking on PIA, these morphologically similar isolates were inoculated into YEPG broth amended with [14 C]triclosan and incubated at 28³C. Both isolates were able to liberate
Fig. 4. Comparison of (A) BOX-PCR, (B) RISA and (C) ARDRA patterns from strain Rd1 and the total consortium DNA. Lane L represent the same ladders used in Fig. 3. R represents strain Rd1 while Cs represent the consortium. Arrows in C represent bands unique to strain Rd1 and the consortium.
FEMSEC 1245 5-7-01
110
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
broth and plate cultivation suggested that strain Rd1 could not use any of these compounds as a sole source of carbon or energy for growth. 3.4. Phylogenetic analysis of consortium members BOX-PCR, RISA and ARDRA ¢ngerprinting of strain Rd1 showed bands that had been present in the consortium pro¢le but that had been missing from any of the other ¢ve isolate pro¢les (Fig. 4A^C). Sequencing of the 16S rRNA gene of strain Rd1 revealed that it was most closely related to a cluster of previously characterized KProteobacteria in the Sphingomonas group and was identical to the Sphingomonas-like partial sequences ampli¢ed and cloned from the consortium lysate. Sequence comparisons revealed that several of the partial 16S rDNA sequences from consortium isolates were 100% identical to the corresponding region of rDNA from previously cultivated organisms or rDNA sequences from uncultivated organism deposited in public databases. The partial rDNA sequences from strains Spd and Ly were each identical to a handful of sequences from bacteria, many of which were involved in the degradation of organic compounds including chlorinated solvents (Table 2). Strain Swo partial rDNA was identical to Alcaligenes sp. strain HR5 [26]. The remaining partial 16S sequences, from isolates Rd1, Sws and Sy, were not 100% identical to any sequence currently found in a public database. The identity of those partial sequences ranged from 98.5 to 99.1% (Table 2). 4. Discussion A stable enrichment culture was obtained that was able to use triclosan as a sole source of carbon and energy. This is the ¢rst report of a bacterial consortium able to use triclosan as a sole source of carbon and energy. Despite numerous attempts to isolate triclosan-degrading strains on solid MSMT, no growth was ever observed on those plates. Contrary to several recent reports [15,27,28], resuspension of cells from the initial dilution plates also failed to yield a single isolate that was capable of degrading triclosan in liquid medium. The initial failure to isolate pure strains from stable
enrichment cultures may indicate that cometabolic steps are required to degrade the substrate [29]. In the case of a recently characterized atrazine-degrading consortium [30], failure to isolate a single strain able to completely mineralize atrazine was due to distribution of the necessary catabolic genes between two di¡erent strains. In other instances, the organism responsible for the metabolic reaction of interest may be fastidious, requiring cofactors from other consortium members [31]. Our inability to cultivate strain Rd1 on minimal media containing either triclosan or other single carbon sources suggests that this strain may fall into this last category, requiring cofactors supplied only in the consortium or complex media. The apparent lag in growth despite immediate mineralization of triclosan (Figs. 1 and 2) suggests that triclosan degradation may be a detoxi¢cation mechanism. This hypothesis is supported by the isolation of Rd1M, a mutant of Rd1 that is impaired in its ability to mineralize triclosan [32]. While Rd1 readily grows on YEPG containing 500 mg l31 triclosan, the growth of strain Rd1M is inhibited by triclosan concentrations in excess of 50 mg l31 . Further work needs to be done to determine what e¡ect degradation has on the overall resistance of Rd1 to triclosan. Despite clear evidence for the partial mineralization of the 14 C-labeled dichloro ring by the consortium and by strain Rd1, no triclosan metabolites were detected by either GC-MS or HPLC analysis of culture supernatants. Previous work by others on the bacterial degradation of various substituted and unsubstituted diphenyl ethers suggests that degradation may occur by either of two mechanisms. One mechanism involves dihydroxylation of one of the rings in the 2,3 positions followed by either intradiol [33] or extradiol [34,35] cleavage. The second mechanism requires a highly regioselective, angular dioxygenation at the 1,2 positions of one of the rings [36], which results in the spontaneous ¢ssion of the ether bond, yielding both a phenol and a catechol moiety [37^40]. Further work is needed to determine whether strain Rd1 relies on either of these, or another, as yet unde¢ned mechanism for the degradation of triclosan. A recent report by Hundt et al. identi¢ed dichlorophenol and several glycosylated triclosan derivatives as transformation products of triclosan by the fungal species Pycnoporus cinnabarinus and Trametes versicolor [41]. Although we were able to detect 2,4-dichlorophenol in
Table 2 Accession numbers and closest 16S match for organisms isolated from the triclosan-degrading consortium Consortium isolate (accession number)
Closest 16S match (accession number)
Ly Spd Swo Sy Sws Rd1
Pseudomonas mendocina Pseudomonas aeruginosa Alcaligenes xylosoxidans Rhodanobacter lindanoclasticus Agrobacterium radiobacter Sphingomonas sanguis
(AF292237) (AF292239) (AF292240) (AF292242) (AF292241) (AF292238)
FEMSEC 1245 5-7-01
Identity (%) (AF232713) (AB037560) (AJ002808) (AF039167) (D137260) (AF226204)
100 100 100 99.1 98.8 98.5
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112
analytical standards down to 100 ng l31 via HPLC, there was no evidence for 2,4-dichlorophenol production either from the consortium or from strain Rd1 individually. When strain Rd1 was grown on complex media, the extent of 14 C-labeled triclosan mineralization was similar to that of the consortium growing on minimal media (data not shown). This suggests that Rd1 is responsible for the bulk of the triclosan mineralization observed and that none of the other consortium members can signi¢cantly degrade the remaining labeled triclosan residues. However, this does not preclude the participation of other consortium members in the degradation of unlabeled, monochloro ring, as total chloride release of greater than 85% indicates signi¢cant transformation of both rings (Table 1). While the partial 16S rDNA sequences obtained from the six isolates are insu¤cient for a detailed phylogenetic determination, it is nevertheless clear that each strain can be assigned to a speci¢c subgroup among the K (strains Rd1 and Sws), L (strain Swo) or Q (strains Spd, Ly, and Sy) Proteobacterial subdivisions. Particularly noteworthy among the isolates is strain Sy, which represents one of only two cultivated members of the Rhodanobacter lindanoclasticus subgroup of the Q-Proteobacteria [42]. R. lindanoclasticus is a novel species that was isolated using the insecticide and wood preservative lindane (Q-hexachlorocyclohexane) as a sole carbon source. Although strain Sy appears to be phylogenetically related to organisms involved in the degradation of chlorinated organic compounds, its role and the role of the other consortium members in the mineralization of triclosan is unclear. Strains from several di¡erent genera have been described which are able to degrade various diphenyl ethers. These genera include Erwinia [43], Pseudomonas [33,35,44], and Sphingomonas [38]. It is interesting to note that only Sphingomonas-like strains have thus far been reported to degrade chlorinated diphenyl ethers [38,39]. Strain Rd1 is the ¢rst organism reported to partially mineralize triclosan. The isolation of strain Rd1 was guided in large part by the sequence information obtained through random cloning and sequencing of partial 16S rRNA gene fragments from the consortium DNA extract. Teske et al. [45] used 16S rDNA sequence information to design selective enrichment conditions for the isolation of individual organisms from a mixed culture. Other molecular-genetic techniques such as ARDRA and REP-PCR have also been used to guide the isolation of refractory organisms from mixed cultures [15]. This work adds to the evidence demonstrating the value of molecular ecology techniques in the characterization of microbial consortia [15,45]. Acknowledgements This work was supported in part by a Dow Foundation SPHERE award to G.S.S. and in part by the Waste Man-
111
agement Research and Education Institute, University of Tennessee, Knoxville. A.G.H. was supported by an appointment to the Alexander Hollaender Distinguished Postdoctoral Fellowship Program sponsored by the US Department of Energy, O¤ce of Health and Environmental Research, and administered by the Oak Ridge Institute for Science and Education. The authors wish to thank Josef Inauen of Ciba Specialty Chemical for the generous gifts of labeled and unlabeled triclosan.
References [1] McMurry, L.M., Oethinger, M. and Levy, S.B. (1998) Triclosan targets lipid synthesis. Nature 394, 531^532. [2] Daughton, C.G. and Ternes, T.A. (1999) Pharmaceuticals and personal care products in the environment : agents of subtle change? Environ. Health Perspect. 107, 907^937. [3] Parikh, S., Xiao, G. and Tonge, P.J. (2000) Inhibition of InhA, the enoyl reductase from mycobacterium tuberculosis, by triclosan and isoniazid. Biochemistry 39, 7645^7650. [4] Heath, R.J. and Rock, C.O. (2000) A triclosan-resistant bacterial enzyme. Nature 406, 145^146. [5] Schweizer, H.P. (1998) Intrinsic resistance to inhibitors of fatty acid biosynthesis in Pseudomonas aeruginosa is due to e¥ux: application of a novel technique for generation of unmarked chromosomal mutations for the study of e¥ux systems. Antimicrob. Agents Chemother. 42, 394^398. [6] Difco Laboratories (1985) Difco Laboratories Incorporated, Detroit, MI. [7] Voets, J.P., Pipyn, P., Van Lancker, P. and Verstraete, W. (1976) Degradation of microbicides under di¡erent environmental conditions. J. Bacteriol. 40, 67^72. [8] Lopez-Avila, V. and Hites, R.A. (1980) Organic compounds in an industrial wastewater : their transport into sediments. Environ. Sci. Technol. 14, 1382^1390. [9] Hale, R.C., Smith, C.L., De Fur, P.O., Harvey, E., Bush, E.O., La Guardia, M.J. and Vadas, G.G. (2000) Nonylphenols in sediments and e¥uents associated with diverse wastewater outfalls. Environ. Toxicol. Chem. 19, 946^952. [10] Okumura, T. and Nishikawa, Y. (1996) Gas chromatography-mass spectrometry determination of trilcosans in water, sediment, and ¢sh samples via methylation with diazomethane. Anal. Chim. Acta 325, 175^184. [11] Miyazaki, T., Yamagishi, T. and Matsumoto, M. (1984) Residues of 4-chloro-1-(2,4-dichlorophenoxy)-2-methoxybenzene (triclosan methyl) in aquatic biota. Bull. Environ. Contam. 32, 227^232. [12] Beck, H.A.D., Eckart, K., Mathar, W. and Wittkowski, R. (1989) Determination of PCDDs and PCDFs in Irgasan DP 300. Chemosphere 19, 167^170. [13] McCullar, M.V., Brenner, V.H.A.R. and Focht, D.D. (1994) Construction of a novel polychlorinated biphenyl-degrading bacterium: Utilization of 3,4P-dichlorobiphenyl by Pseudomonas acidovorans M3GY. Appl. Environ. Microbiol. 60, 3833^3839. [14] Arensdorf, J.J. and Focht, D.D. (1995) A meta cleavage pathway for 4-chlorobenzoate, an intermediate in the metabolism of 4-chlorobiphenyl by Pseudomonas cepacia P166. Appl. Environ. Microbiol. 61, 443^447. [15] Maltseva, O. and Oriel, P. (1997) Monitoring of an alkaline 2,4,6trichlorophenol-degrading enrichment culture by DNA ¢ngerprinting methods and isolation of the responsible organism, haloalkiphilic Nocardioides sp. strain M6. Appl. Environ. Microbiol. 63, 4145^ 4149. [16] Hay, A.G. and Focht, D.D. (1998) Cometabolism of 1,1-dichloro-
FEMSEC 1245 5-7-01
112
[17]
[18]
[19]
[20]
[21]
[22] [23]
[24]
[25]
[26]
[27]
[28]
[29]
[30]
A.G. Hay et al. / FEMS Microbiology Ecology 36 (2001) 105^112 2,2-bis(4-chlorophenyl)ethylene by Pseudomonas acidovorans M3GY grown on biphenyl. Appl. Environ. Microbiol. 64, 2141^2146. Focht, D.D. and Alexander, M. (1971) Aerobic cometabolism of DDT analogues by Hydrogenomonas sp.. J. Agric. Food Chem. 19, 20^22. Ausubel, F.M., Brent, R., Kingston, R.E., More, D.D., Seidman, J.G., Smith, J.A. and Struhl, K. (1989) Current protocols in Molecular Biology John Wiley and Sons, New York, NY. Rademaker, J.L.W., Louws, F.J. and De Bruijn, F.J. (1998) Characterization of the diversity of ecologically important microbes by repPRC genomic ¢ngerprinting in: Molecular Microbiology Ecology Manual, Vol. 3 (Akkermans, A.D.L., Van Elsas, J.D. and De Bruijn, F.J., Eds.). Kluwer Academic, Dordrecht. Ferris, M.J., Muyzer, G. and Ward, D.M. (1996) Denaturing gradient gel electrophoresis pro¢les of 16S rRNA-de¢ned populations inhabiting a hot spring microbial mat community. Appl. Environ. Microbiol. 62, 340^346. Borneman, J. and Triplett, E.W. (1997) Molecular microbial diversity in soils from eastern Amazonia: Evidence for unusual microorganisms and microbial population shifts associated with deforestation. Appl. Environ. Microbiol. 63, 2647^2653. Roux, K.H. (1994) Using mismatched primer-template pairs in touchdown PCR. BioTechniques 16, 812^814. Vaneechoutte, M. et al. (1992) Rapid identi¢cation of bacteria of the Comamonadaceae with ampli¢ed ribosomal DNA-restriction analysis (ARDRA). FEMS Microbiol. Lett. 93, 227^234. Lane, D.J. (1991) 16S/235 rRNA sequencing in: Nucleic Acid Techniques in Bacterial Systematics (Stackebrandt, E. and Goodfellow, M., Eds.), pp. 115^147. John Wiley and Sons, New York. Altschul, S.F., Madden, T.L., Scha¡er, A.A., Zhang, J., Zhang, Z., Miller, W. and Lipman, D.J. (1997) Gapped BLAST and PSIBLAST: a new generation of protein database search programs. Nucleic Acids Res. 25, 3389^3402. Ho¡mann, A., Thimm, T., Droege, M., Moore, E.R.B., Munch, J.C. and Tebbe, C.C. (1998) Intergeneric transfer of conjugative and mobilizable plasmids harbored by Escherichia coli in the gut of the soil microarthropod Folsomia candida (Collembola). Appl. Environ. Microbiol. 64, 2652^2659. Sutherland, T.D., Horne, I., Lacey, M.J., Harcourt, R.L., Russel, R.J. and Oakeshott, J.G. (2000) Enrichment of an endosulfane-degrading mixed bacterial culture. Appl. Environ. Microbiol. 66, 2822^ 2828. Tanghe, T., Dhooge, W. and Verstraete, W. (1999) Isolation of a bacterial strain able to degrade branched nonylphenol. Appl. Environ. Microbiol. 65, 746^751. Focht, D.D. (1993) in: Soil Biochemistry, Vol. 8 (Bollag, M. and Stozky, G., Eds.). Marcel Dekker, New York Microbial degradation of chlorimated biphenyls, 341^407. De Souza, M.L., Newcombe, D., Alvey, S., Crowley, D., Hay, A., Sadowsky, M.J. and Wackett, L.P. (1998) Molecular basis of a bac-
[31]
[32] [33]
[34]
[35]
[36]
[37]
[38]
[39]
[40]
[41]
[42] [43]
[44]
[45]
terial consortium: interspecies catabolism of atrazine. Appl. Environ. Microbiol. 64, 178^184. Maymo-Gatell, X., Chien, Y.-T., Gossett, J.M. and Zinder, S.H. (1997) Isolation of a bacterium that reductively dechlorinates tetrachloroethylene to ethene. Science 276, 1568^1571. Hay, A.G., Unpublished results. Takase, I., Omori, T. and Minoda, Y. (1986) Microbial degradation products from biphenyl-related compounds. Agric. Biol. Chem. 50, 681^686. Pfeifer, F., Trueper, H.-G., Klein, J. and Schacht, S. (1993) Degradation of diphenylether by Pseudomonas cepacia Et4: Enzymatic release of phenol from 2,3-dihydroxydiphenyl ether. Arch. Microbiol. 159, 323^329. Pfeifer, F., Schacht, S., Klein, J. and Truper, H.G. (1989) Degradation of diphenylether by Pseudomonas cepacia. Arch. Microbiol. 152, 515^519. Fortnagel, P., Wittich, R.M., Harms, H., Schmidt, S., Franke, S., Sinnwell, V., Wilkes, H. and Francke, W. (1989) New bacterial degradation of the biaryl ether structure regioselective dioxygenation prompts cleavage of ether bonds. Naturwissenschaften 76, 523^524. Schmidt, S., Wittich, R.M., Fortnagel, P., Erdmann, D. and Francke, W. (1992) Metabolism of 3-methyldiphenyl ether by Sphingomonas sp. SS31. FEMS Microbiol. Lett. 96, 253^258. Schmidt, S., Wittich, R.M., Erdmann, D., Wilkes, H., Francke, W. and Fortnagel, P. (1992) Biodegradation of diphenyl ether and its monohalogenated derivatives by Sphingomonas sp. strain SS3. Appl. Environ. Microbiol. 58, 2744^2750. Schmidt, S., Fortnagel, P. and Wittich, R.M. (1993) Biodegradation and transformation of 4,4P- and 2,4-dihalodiphenyl ethers by Sphingomonas sp. strain SS33. Appl. Environ. Microbiol. 59, 3931^3933. Wilkes, H., Francke, W., Wittich, R.M., Harms, H., Schmidt, S. and Fortnagel, P. (1992) Mechanistic investigations on microbial degradation of diaryl ethers analysis of isotope-labeled reaction products. Naturwissenschaften 79, 269^271. Hundt, K., Martin, D., Hammer, E., Jonas, U., Kindermann, M.K. and Schauer, F. (2000) Transformation of triclosan by Trametes versicolor and Pycnoporus cinnabarinus. Appl. Environ. Microbiol. 66, 4157^4160. Maidak, B.L. et al. (2000) The RDP (Ribosomal Database Project) continues. Nucleic Acids Res. 28, 173^174. Liaw, J.H. and Srinivasan (1989) Molecular cloning and expression of an Erwinia sp. gene encoding diphenyl ether cleavage in Escherichia coli. Appl. Environ. Microbiol. 55, 2220^2225. Wittich, R.M., Schmidt, S. and Fortnagel, P. (1990) Bacterial degradation of 3- and 4-carboxybiphenyl ether by Pseudomonas sp. NSS2. FEMS Microbiol. Lett. 67, 157^160. Teske, A.P.S., Cohen, Y. and Muyzer, G. (1996) Molecular identi¢cation of bacteria from a coculture by denaturing gradient gel electrophorsis of 16S ribosomal DNA fragments as a tool for isolation in pure cultures. Appl. Environ. Microbiol. 62, 4210^4215.
FEMSEC 1245 5-7-01