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1 LEPAE – Departamento de Engenharia Química, Faculdade de Engenharia, Universidade do Porto, 4200-465 Porto, Portugal
2 Escola Superior de Biotecnologia, Universidade Católica Portuguesa, 4200-072 Porto, Portugal
3 Instituto de Tecnologia Química e Biológica, Av. da República - EAN, 2780-157 Oeiras, Portugal
Correspondence
Olga C. Nunes
opnunes{at}fe.up.pt
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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; Kato et al., 2005). External perturbations, such as the introduction of large quantities of xenobiotics into the environment, create strong selective pressures. One expected reaction to such a stimulus is the establishment of communities able to degrade these compounds. The elucidation of the contribution of individual members of a microbial community to metabolic degradative pathways may shed light on the biochemical transformations taking place and on the interrelationships responsible for the stable coexistence of the organisms. Several studies reporting co-operation among consortium members during the degradation of pollutants have shown different modes of microbial interaction. The literature available shows that the distribution of metabolic steps among consortium members may rely upon different forms of association, namely cross-feeding (e.g. De Souza et al., 1998; Dejonghe et al., 2003), detoxification (e.g. Feigel & Knackmuss, 1993; Pelz et al., 1999) or supply of nutritional requirements (e.g. Hay et al., 2001; Sorensen et al., 2002).
The study of the evolution of degradative pathways has been another area of interest, encouraged by the need to deal with environmental contamination with xenobiotics. Some evidence suggests that an original metabolic association of various micro-organisms may evolve to a single-organism-based process (e.g. De Souza et al., 1998). Such a transformation can be explained on the basis of horizontal gene transfer, leading to the assembly of different catabolic mobile genetic elements in a single cell. The origination of new catabolic routes by the rearrangement and combination of pre-existing genes of different micro-organisms has been reported (Tsuda et al., 1999; Top et al., 2002; Nojiri et al., 2004). Thus, the description of metabolic pathways before the occurrence of horizontal transfer of catabolic genes may provide insights into the evolution of microbial degradation of xenobiotic compounds.
In a previous report (Barreiros et al., 2003) we showed that a defined mixed culture composed of five bacterial isolates (mixed culture DC) could mineralize the herbicide molinate without the accumulation of degradation products. Isolates ON1 and ON3 were identified as Pseudomonas chlororaphis and Pseudomonas nitroreducens, respectively, isolate ON2 as Stenotrophomonas maltophilia and isolate ON5 as Achromobacter xylosoxidans; isolate ON4, not affiliated to any validly named taxon, represented a new genus and species, Gulosibacter molinativorax, within the family Microbacteriaceae (Manaia et al., 2004). In mixed culture DC, G. molinativorax ON4T was able to degrade molinate into ethanethiol and another compound that could not be detected but that was presumed to be an azepane derivative of the herbicide (Barreiros et al., 2003). In spite of this degradative activity, G. molinativorax ON4T was not able to grow in axenic culture with molinate at concentrations above 2 mM, an effect that was abolished when the Gram-negative members of the mixed culture were grown in a vial placed in the headspace of the growth vessel. This observation, along with the monitoring of sulphur compounds (ethanethiol and diethyl disulphide) formed in the headspace of the axenic but not in the mixed culture, suggested a detoxifying association between G. molinativorax ON4T and its commensals. In this study, the fate of the azepane derivative and the role of each of the five members of culture DC in molinate mineralization were further investigated.
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METHODS |
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Growth media and culture conditions.
Culture DC and isolates ON1–ON5 were grown, in axenic or defined mixed cultures, in mineral medium B (Barreiros et al., 2003) supplemented with molinate or other substrate, at 30 °C and 120 r.p.m. Inocula of culture DC were grown in medium B with 4 mM molinate and those of axenic cultures were grown on Luria–Bertani medium (LB) with agar (2 % w/v), supplemented with 1 mM molinate (modified LA) to maintain the selective pressure in the medium.
The anaerobic degradation of molinate by G. molinativorax ON4T and by culture DC was assayed in 100 ml PTFE-sealed flasks containing 75 ml medium B supplemented with 1 mM molinate and 0.01 % (w/v) potassium nitrate, and incubated under a nitrogen atmosphere for 15 days. Samples were collected over time and were analysed for biomass, molinate, dissolved organic carbon (DOC) and ethanethiol contents.
The ability of each member of culture DC to use the azepane moiety derivative azepane-1-carboxylic acid (ACA) resulting from molinate degradation was assessed by incubating each individual organism in the cell-free liquid phase of anaerobic resting cells of G. molinativorax ON4T. ACA was produced by incubating resting cells of G. molinativorax ON4T in sterile saline solution supplemented with 2 mM molinate, under a nitrogen atmosphere, at 30 °C and 120 r.p.m. for 12 h, as described below. The suspension was centrifuged (15 500 g, 10 min); the supernatant was filter-sterilized (0.2 µm pore size) and supplemented with medium B components and magnesium sulphate (0.4 mM) as a sulphur source. Growth was monitored over 7 days.
The ability of culture DC and its members to use different substrates as carbon and nitrogen sources was tested in medium B supplemented with 0.4 mM magnesium sulphate and 5.5 mM HMI, caprolactam or 6-aminohexanoic acid. When necessary, the medium was supplemented with 4 mM ammonium sulphate (as nitrogen source) or with 0.2 g yeast extract l–1 (as source of growth factors). Cultures were incubated aerobically in 100 ml screw-capped Erlenmeyer flasks with Teflon-lined caps. Cell growth was monitored over 7 days.
To compare the molinate mineralization efficiency of culture DC and of the mixture of P. chlororaphis ON1 and G. molinativorax ON4T, these cultures were grown aerobically in 100 ml PTFE-sealed flasks containing 20 ml medium B with 4 mM molinate. Culture samples were collected at regular intervals and analysed for biomass, molinate and DOC contents. The presence of molinate S-ethyl and azepane moieties was also analysed, in the headspace (SPME-GC-FID) and in the liquid phase of cultures (NMR), respectively. An axenic culture of G. molinativorax ON4T and uninoculated medium were analysed similarly, as controls.
To assess the stability of mixed culture DC, cells were grown in medium B with 4 mM molinate for 5 days at 30 °C, stored at 4 °C for 8–20 days, transferred to fresh medium and incubated for a further 5 days at 30 °C. Culture composition was assessed by PCR-denaturing gradient gel electrophoresis (DGGE) after a total of 20 successive culture transfers.
Resting cell assays.
The isolates were grown on modified LA medium at 30 °C, for 1 or 3 days, centrifuged, washed and resuspended in phosphate buffer 54 mM, pH 7.2 (PB), supplemented with the substrate to test. For NMR analysis, PB was replaced by 50 mM NaCl. Cell densities corresponding to 4 g cell dry weight l–1 were used in all assays. To evaluate the ability of each isolate to consume ethanethiol, resting cell assays were performed in PB supplemented with 0.4 mM of this sulphur compound. Resting cells of culture DC were used as positive control and uninoculated buffer with ethanethiol was used to assess abiotic losses. Sulphur compound content was measured in the headspace (SPME-GC-FID). To obtain and/or identify the azepane derivative, resting cells of G. molinativorax ON4T were incubated in saline solution supplemented with 2 mM molinate under a nitrogen atmosphere, at 30 °C and 120 r.p.m. Samples were collected over time and the liquid phase was analysed for molinate content (HPLC) or for detection and identification of molinate metabolites (NMR).
To evaluate if the use of HMI and caprolactam by G. molinativorax ON4T were induced processes, resting cells pregrown with 2 mM of the azepane moiety derivative of molinate (ACA) were incubated in PB with 2 mM HMI and/or caprolactam. Abiotic losses were assessed in uninoculated buffer with the same concentration of HMI and/or caprolactam. Samples were collected over time and analysed by NMR or HPLC to follow HMI or caprolactam degradation, respectively. Pseudomonas strains ON1 and ON3 were also tested for ACA-HMI induction of caprolactam degradation, using the same procedure.
Analytical procedures.
Cell growth was monitored spectrophotometrically (OD610; Philips PU-8620UV/VIS spectrophotometer) and cell dry weight was obtained via a calibration curve (Barreiros et al., 2003). Molinate and DOC contents in culture supernatants were analysed (HPLC and total organic carbon) as described before (Barreiros et al., 2003). Ethanethiol was quantified in the liquid phase using the colorimetric method described by Dias & Weimer (1998). The analysis of ethanethiol, diethyl disulphide and ethyl methyl sulphide in the headspace of cultures or resting cells assays was performed by SPME-GC-FID as described by Barreiros et al. (2003).
NMR analysis.
Proton spectra were recorded on Bruker DRX500 or AMX300 spectrometers operating, respectively, at 500.13 MHz and 300.13 MHz for 1H, equipped with 5 mm diameter inverse detection broadband probe heads (with and without XYZ gradients). In both cases water pre-saturation was applied. Acquisition parameters on DRX500: spectral width, 10 kHz; pulse width, 7 ms (6 ° flip angle); data points, 64K; repetition delay, 1 s; number of transients, 100. Acquisition parameters on AMX300: spectral width, 5 kHz; pulse width, 4 ms (7 ° flip angle); data points, 16K; repetition delay, 1.5 s; number of transients, 100. Carbon spectra were recorded on a Bruker DRX500 operating at 125.77 MHz for 13C and equipped with a 1H/13C dual-probe head (5 mm diameter). Acquisition parameters: spectral width, 31.5 kHz; pulse width, 6 ms (7 ° flip angle); data points, 64K; repetition delay, 10 s; number of transients, 5500. Proton broadband decoupling was applied during acquisition time only (1.04 s). All the spectra were acquired at a constant temperature of 27 °C. Chemical shifts were referenced to tetramethylsilane at 0.0 p.p.m. (1H) and methanol at 49.3 p.p.m. (13C), used as external references.
Denaturing gradient gel electrophoresis.
Total genomic DNA was extracted based on the method described by Cashion et al. (1977) from 0.35 mg biomass of culture DC or of each isolate. A 200 bp fragment (based on the reference strain Escherichia coli bases 338 and 518) of the 16S rRNA gene was amplified using the bacterial 16S rDNA primers forward 338F_GC, containing a GC clamp (5'-CGC CCG CCG CGC GCG GCG GGC GGG GCG GGG GCA CGG GGG GAC TCC TAC GGG AGG CAG CAG-3'), and reverse 518R (5'-ATT ACC GCG GCT GCT GG-3') (Muyzer et al., 1993). Reaction mixtures (50 µl) were prepared as described by Henriques et al. (2006). The PCR conditions were: 5 min at 94 °C, 30 cycles of (30 s at 92 °C, 30 s at 55 °C, 30 s at 72 °C), and 7 min at 72 °C. PCR products were separated in a 8 % (w/v) polyacrylamide gel with a denaturing gradient ranging from 30 to 55 % (where 100 % denaturant contained 7 M urea and 40 % formamide), running initially at 20 V for 20 min, and then at 200 V for 5.5 h at 60 °C. DGGE bands corresponding to each culture DC isolate were excised from the gel, eluted with 20 µl ultrapure water, reamplified by PCR with the same primers, cleaned (GFX PCR DNA and Gel Band Purification kit, Amersham Biosciences) and subsequently sequenced with primer 518R. The sequences were determined using a model ABI 3700 DNA analyser (Applied Biosystems), and their quality was checked manually using the BioEdit software (Hall, 1999). These sequences were compared with those obtained for the 16S rRNA gene of each isolate (accession numbers: P. chlororaphis ON1, AJ306832; S. maltophilia ON2, AJ306833; P. nitroreducens ON3, AJ306834; G. molinativorax ON4T, AJ306835; A. xylosoxidans ON5, AJ306836).
Kinetic models.
Specific cell growth and molinate degradation rates were calculated during the exponential growth phase according to the first-order kinetic model described by Barreiros et al. (2003). The same data were fitted to the Haldane equation:
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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). However, the limitation of the growth conditions and analytical methods (HPLC and GC-MS) used could also hamper the detection of the azepane moiety. In fact, it was observed that under anaerobic conditions mixed culture DC degraded 90 % (w/w) of the added molinate, whereas only 28 % of the dissolved organic carbon (DOC) was consumed after 80 h of incubation (Correia et al., 2006). These data suggested the accumulation of the molinate azepane moiety, although it could not be detected by HPLC or GC-MS. Therefore, it was hypothesized that, in the absence of oxygen, the hydrolysis of the molinate thioester bond could occur as long as G. molinativorax ON4T was viable, although the further degradation of the metabolites was not possible. This hypothesis was tested by culturing mixed culture DC and G. molinativorax ON4T anaerobically. Under this condition, mixed culture DC was not able to grow, although the herbicide, at an initial concentration of 
160 mg l–1, was slowly degraded to concentrations below the detection limit (0.9 mg l–1). Ethanethiol, a major product of molinate breakdown by G. molinativorax ON4T, was not detected in the medium. In spite of this, after 15 days of incubation the initial DOC content was reduced by only 30 % (Table 1). In a parallel experiment, G. molinativorax ON4T showed different results: only 45 % (w/w) of the molinate was degraded; ethanethiol accumulated in the medium (18 mg l–1 in the liquid phase), and the initial DOC content was reduced by only 12 % (Table 1). These final DOC values were consistent with those that would result from partial molinate hydrolysis. These results suggest that under anaerobic conditions G. molinativorax ON4T maintains its ability to break down the molinate thioester bond. Apparently, none of the mixed culture DC members is able to further metabolize the azepane derivative that accumulates in the medium. In contrast, ethanethiol was consumed by the Gram-negative isolates of mixed culture DC. Thus, the toxic effect of that thiol on G. molinativorax ON4T was attenuated and it was able to proceed with molinate hydrolysis.
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). During incubation, the resonances assigned to molinate (–S–CH2CH3 at 1.20 p.p.m.; –S–CH2CH3 at 2.82 p.p.m.; azepane ring at 3.47, 1.70, 1.63 and 1.48 p.p.m.) decreased and the formation of a product attributed to the molinate azepane moiety was observed. Small amounts of free ethanethiol remained in solution and were detected (HS-CH2CH3 at 1.12 p.p.m.; HS-CH2CH3 at 3.60 p.p.m.). After 6 h incubation, all the molinate was converted into degradation products that were not further modified, even after 30 h incubation. This procedure proved to be a good process to obtain high amounts of the azepane-containing compound and, hence, to proceed with its characterization by 1H and 13C NMR (Fig. 2a, b). 1H and 13C-NMR chemical shifts characteristic of the new compound, identified as azepane-1-carboxylic acid (ACA), observed in saline solution (50 mM NaCl) at physiological pH are indicated in Table 2. In Fig. 2(a), resonances in the proton spectrum are shifted to the low-field region due to ionic strength of the solution (approx. 5 M NaCl) upon lyophilization of the liquid phase of anaerobic resting cells of G. molinativorax ON4T with molinate after 12 h incubation. This is believed to be the first report of ACA as an intermediate of molinate degradation, suggesting that G. molinativorax ON4T may use a unique process to degrade this herbicide.
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McClung et al. (1994) described a microbiological process of degradation of 1-ethylsulphanyl-N,N-dipropylformamide (EPTC), which consisted of the hydrolysis of the thioester bond, with release of ethanethiol, dipropylamine and CO2. Hypothesizing an equivalent transformation for molinate, the organisms able to use ACA – Pseudomonas strains ON1 and ON3 and G. molinativorax ON4T – would promote its decarboxylation into hexamethyleneimine (HMI), which could be further consumed as a source of carbon and nitrogen. Such a supposition was consistent with previous reports on molinate biodegradation that refer to the release of the azepane moiety in the form of HMI as a degradation product of the intermediate molinate sulphoxide (Thomas & Holt, 1980; Imai & Kuwatsuka, 1986). In order to confirm the hypothesis that further molinate degradation could occur via HMI, this compound, and other commercially available possible analogues of the degradation metabolites of ACA, were assayed as growth substrates. In fact, all the isolates able to degrade ACA were also able to grow at the expense of HMI. However, in the Pseudomonas strains this process was constitutive, whereas in G. molinativorax ON4T it was subject to ACA induction. These data suggest that ACA decarboxylation to HMI, by analogy with the metabolic pathway proposed by McClung et al. (1994), represents the next step of molinate mineralization. According to Wackett & Hershberger (2001), HMI degradation may follow one of two alternative pathways, via caprolactam or via an open-ring aldehyde metabolite. Pseudomonas strains ON1 and ON3 were unable to grow at the expense of caprolactam, even after induction with HMI. Thus, HMI degradation in these strains may proceed via 2-hydroxy-HMI, followed by isomerization into an open-ring aldehyde metabolite and further oxidization to 6-aminohexanoic acid. In contrast, caprolactam was degraded by G. molinativorax ON4T, although only after successive induction with ACA and HMI. This finding hints that HMI degradation in G. molinativorax ON4T occurs via a distinct pathway, where caprolactam is formed and further converted into 6-aminohexanoic acid. This last compound, common to both HMI degradation routes, supported the growth of both pseudomonads and of G. molinativorax ON4T.
Degradation of the S-ethyl moiety of molinate
Ethanethiol and diethyl disulphide, formed during the initial breakdown of molinate, are toxic to G. molinativorax ON4T. However, in mixed culture DC this toxic effect is avoided because P. chlororaphis ON1 and S. maltophilia ON2 are able to consume these sulphur compounds (Barreiros et al., 2003). Further investigation of the degradative performance of these two isolates revealed that P. chlororaphis ON1 consumed ethanethiol and diethyl disulphide at a higher rate than S. maltophilia ON2 (Table 3). These results were confirmed by the specific degradation rates of resting cells of these isolates with 1 mM ethanethiol. P. chlororaphis ON1 degraded this compound at a rate approximately 10 times higher [0.36±0.13 mg ethanethiol (g cell dry weight)–1 h–1] than S. maltophilia ON2 [0.03±0.009 mg ethanethiol (g cell dry weight)–1 h–1]. P. nitroreducens ON3 was also able to transform ethanethiol, although it was observed to be a conversion only, as no depletion of the added carbon occurred. In fact, approximately 75 % of the added ethanethiol was converted to ethyl methyl sulphide. Given that none of the culture DC members was able to convert or to degrade the ethyl methyl sulphide, and considering the high rates of ethanethiol transformation during molinate mineralization (Barreiros et al., 2003), this product is probably not formed by P. nitroreducens ON3, when S. maltophilia ON2, and particularly P. chlororaphis ON1, are also present. Taken together, these results suggest that, in mixed culture DC, the ethanethiol formed during molinate cleavage can be consumed by P. chlororaphis ON1 and S. maltophilia ON2, although the first, as a more rapid degrader, may compete for this substrate.
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shows a putative degradation pathway for molinate and the flow of carbon, nitrogen and sulphur through the different members of culture DC, as suggested by the results obtained in the present study. According to this proposed pathway, molinate can be mineralized by G. molinativorax ON4T and P. chlororaphis ON1. First, molinate is hydrolysed into ethanethiol and ACA by G. molinativorax ON4T, then ethanethiol is consumed by P. chlororaphis ON1, whereas ACA supports the growth of both strains. The behaviour of the mixture of these isolates in the presence of 4 mM molinate was compared with that of mixed culture DC. The analysis of Fig. 4(a, b) shows that both mixed cultures were able to grow on molinate with concomitant DOC depletion to values of 20–30 mg l–1. SPME-GC-FID analysis of the culture headspace and 1H-NMR of the respective supernatant after 72 h incubation revealed that neither the S-ethyl moiety (ethanethiol and diethyl disulphide) nor ACA accumulated in the medium (results not shown). However, the comparison of the specific growth and molinate degradation rates of the two mixed cultures revealed that, although molinate was mineralized by the mixture of P. chlororaphis ON1 and G. molinativorax ON4T, a possible substrate inhibition may occur. While the experimental data of growth and molinate degradation by culture DC fit a first-order kinetics [0.06 h–1 and 208 mg molinate (g cell dry weight)–1 h–1, respectively], as described before (Barreiros et al., 2003), the data corresponding to the mixture of P. chlororaphis ON1 and G. molinativorax ON4T could be described by the Haldane equation (see Methods), a conventional model of substrate inhibition. The Haldane model parameters obtained for culture DC and the mixture of P. chlororaphis ON1 and G. molinativorax ON4T (with Ks=1 mg l–1) were, respectively, µmax 0.38 h–1 and 1.39 h–1, and Ki 102.0 mg l–1 and 11.5 mg l–1. The specific growth rate of the mixture of P. chlororaphis ON1 and G. molinativorax ON4T increases from 0.026 h–1 for initial molinate concentration of about 600 mg l–1 up to 0.04 h–1 after consumption of approximately half the initial molinate concentration.
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), as confirmed by the comparison of the sequences of each excised band with those of the pure isolates of mixed culture DC.
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reported the stable co-existence of a member with no apparent function in a community in a chemostat fed with 4-chlorosalicylate. The stability of consortia for long periods may be based upon a complex set of microbial interrelationships, which may include synergistic interactions involving carbon sharing, consumption of toxic metabolites, commensalism and/or competition (e.g. Pelz et al., 1999; Christensen et al., 2002; Dejonghe et al., 2003; Kato et al., 2005).
The observation that micro-organisms have evolved high catabolic potential for numerous pollutants has led to their intensive isolation and characterization, with the ultimate objective of establishing cost-effective bioremediation processes. The accumulation of knowledge on the metabolites/catalysts/genes involved in the breakdown of different organic functional groups permitted the conclusion that different micro-organisms would degrade different compounds of the same class of chemical structure by similar catabolic routes. This allowed for creation of databases of degradative pathways (e.g. University of Minnesota Biocatalysis/Biodegradation Database, http://umbbd.ahc.umn.edu/) of great utility for previewing the catabolic routes of new isolates and/or pollutants. However, possibly due to the evolutionary potential and genetic flexibility of micro-organisms (Timmis & Pieper, 1999), there are still descriptions of novel pathways for the degradation of pollutants for which other catabolic routes were previously known (e.g. Pieper et al., 2004). Apparently this is the case for molinate. Initially it was described as co-metabolizable by three possible degradative routes: (1) the oxidation of the sulphur atom with production of molinate sulphoxide and molinate sulphone; (2) the oxidation of the azepane moiety with production of hydroxy- and oxomolinate; or (3) the oxidation of the S-ethyl moiety with production of molinate alcohol and molinate acid (Soderquist et al., 1977; Thomas & Holt, 1980; Golovleva et al., 1981; Imai & Kuwatsuka, 1986). Among those compounds, only trace amounts of 2-oxomolinate were detected in culture DC grown with molinate as the only source of carbon, nitrogen and energy (Correia et al., 2006), supporting the hypothesis that the herbicide is degraded by this mixed culture through a different catabolic route. We (Barreiros et al., 2003; the present work) have described a new mineralizing pathway, involving the hydrolysis of the herbicide thioester bond by G. molinativorax ON4T with the release of ethanethiol and ACA, which is apparently further degraded, via previously described mechanisms, by Pseudomonas strains ON1 and ON3 and G. molinativorax ON4T.
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ACKNOWLEDGEMENTS |
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Edited by: H. L. Drake
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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Cashion, P., Holder-Franklin, M. A., McCully, J. & Franklin, M. (1977). A rapid method for the base ratio determination of bacterial DNA. Anal Biochem 81, 461–466.[CrossRef][Medline]
Christensen, B. B., Haagensen, J. A., Heydorn, A. & Molin, S. (2002). Metabolic commensalism and competition in a two-species microbial consortium. Appl Environ Microbiol 68, 2495–2502.
Correia, P., Boaventura, R. A., Reis, M. A. & Nunes, O. C. (2006). Effect of operating parameters on molinate biodegradation. Water Res 40, 331–340.[Medline]
De Souza, M. L., Newcombe, D., Alvey, S., Crowley, D. E., Hay, A., Sadowsky, M. J. & Wackett, L. P. (1998). Molecular basis of a bacterial consortium: interspecies catabolism of atrazine. Appl Environ Microbiol 64, 178–184.
Dejonghe, W., Berteloot, E., Goris, J., Boon, N., Crul, K., Maertens, S., Höfte, M., De Vos, P., Verstraete, W. & Top, E. M. (2003). Synergistic degradation of linuron by a bacterial consortium and isolation of a single linuron-degrading Variovorax strain. Appl Environ Microbiol 69, 1532–1541.
Dias, B. & Weimer, B. (1998). Purification and characterization of L-methionine 
-lyase from Brevibacterium linens BL2. Appl Environ Microbiol 64, 3327–3331.
Feigel, B. J. & Knackmuss, H. J. (1993). Syntrophic interactions during degradation of 4-aminobenzenesulfonic acid by a two-species bacterial culture. Arch Microbiol 159, 124–130.[CrossRef][Medline]
Golovleva, L. A., Finkelstein, Z. I., Popovich, N. A. & Skriabin, G. K. (1981). Transformation of ordram by microorganisms. Izv Akad Nauk SSSR Biol 3, 348–358. (in Russian)
Hall, T. A. (1999). BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp Ser 41, 95–98.
Hay, A. G., Dees, P. M. & Sayler, G. S. (2001). Growth of a bacterial consortium on triclosan. FEMS Microbiol Ecol 36, 105–112.[CrossRef][Medline]
Henriques, I. S., Alves, A., Tacão, M., Almeida, A., Cunha, A. & Correia, A. (2006). Seasonal and spatial variability of free-living bacterial community composition along an estuarine gradient (Ria de Aveiro, Portugal). Estuar Coast Shelf Sci 68, 139–148.[CrossRef]
Imai, Y. & Kuwatsuka, S. (1986). Metabolic pathways of the herbicide molinate in four strains of isolated soil microorganisms. J Pestic Sci 11, 245–251.
Kato, S., Haruta, S., Cui, Z. J., Ishii, M. & Igarashi, Y. (2005). Stable coexistence of five bacterial strains as a cellulose-degrading community. Appl Environ Microbiol 71, 7099–7106.
Manaia, C. M., Nogales, B., Weiss, N. & Nunes, O. C. (2004). Gulosibacter molinativorax gen. nov., sp. nov., a molinate degrading bacterium, and classification of ‘Brevibacterium helvolum’ DSM 20419 as Pseudoclavibacter helvolus gen. nov., sp. nov. Int J Syst Evol Microbiol 54, 783–789.
McClung, G., Dick, W. A. & Karns, J. (1994). EPTC degradation by isolated soil microorganisms. J Agric Food Chem 42, 2926–2931.[CrossRef]
Muyzer, G., de Waal, E. C. & Uitterlinden, A. G. (1993). Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol 59, 695–700.
Nojiri, H., Shintani, M. & Omori, T. (2004). Divergence of mobile genetic elements involved in the distribution of xenobiotic-catabolic capacity. Appl Microbiol Biotechnol 64, 154–174.[CrossRef][Medline]
Pelz, O., Tesar, M., Wittich, R. M., Moore, E. R. B., Timmis, K. N. & Abraham, W. R. (1999). Towards elucidation of microbial community metabolic pathways: unravelling the network of carbon sharing in a pollutant-degrading bacterial consortium by immunocapture and isotopic ratio mass spectrometry. Environ Microbiol 1, 167–174.[CrossRef][Medline]
Pieper, D. H., Martins dos Santos, V. A. P. & Golyshin, P. N. (2004). Genomic and mechanistic insights into the biodegradation of organic pollutants. Curr Opin Biotechnol 15, 215–224.[CrossRef][Medline]
Soderquist, C. J., Bowers, J. B. & Crosby, D. G. (1977). Dissipation of molinate in a rice field. J Agric Food Chem 25, 940–945.[CrossRef]
Sorensen, S. R., Ronen, Z. & Aamand, J. (2002). Growth in coculture stimulates metabolism of the phenylurea herbicide isoproturon by Sphingomonas sp. strain SRS2. Appl Environ Microbiol 68, 3478–3485.
Thomas, V. M. & Holt, C. L. (1980). The degradation of [14C]molinate in soil under flooded and nonflooded conditions. J Environ Sci Health B 15, 475–484.[Medline]
Timmis, K. N. & Pieper, D. H. (1999). Bacteria designed for bioremediation. Trends Biotechnol 17, 200–204.[CrossRef][Medline]
Top, E. M., Springael, D. & Boon, N. (2002). Catabolic mobile genetic elements and their potential use in bioaugmentation of polluted soils and waters. FEMS Microbiol Ecol 42, 199–208.[CrossRef]
Tsuda, M., Tan, H. M., Nishi, A. & Furukawa, K. (1999). Mobile catabolic genes in bacteria. J Biosci Bioeng 87, 401–410.[CrossRef][Medline]
Wackett, L. P. & Hershberger, C. D. (2001). Metabolic Logic and Pathway maps. In Biocatalysis and Biodegradation: Microbial Transformation of Organic Compounds, pp. 135–155. Washington, DC: American Society for Microbiology.
Received 22 November 2007;
revised 30 December 2007;
accepted 2 January 2008.
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, Cell dry weight; 
, molinate; 
, DOC; 
, molinate in uninoculated medium. Fittings of a first-order kinetic model for cell dry weight and molinate are shown by full lines and bold full lines, respectively; fittings of the Haldane model for cell dry weight and molinate are shown by dashed lines and bold dashed lines, respectively.