strain FP1

Syed A. Hasan1, Hjalmar P. Permentier1, Anouk Duke2, Carvalho M. Fatima2, Jan van Leeuwen1, Paula M. Castro2, and Dick B. Janssen1

1Department of Biochemistry, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Nijenborgh 4, 9747 AG Groningen,

The Netherlands.

2Escola Superior de Biotecnologia – Universidade Católica Portuguesa, Rua António Bernardino de Almeida

4200-072 Porto, Portugal.

Under review

ABSTRACT

A bacterial strain capable of utilizing 2-fluorophenol as the sole carbon source was isolated from a site contaminated with halogenated aromatic compounds, and identified as a Rhodococcus strain. A combination of proteomic and biochemical experiments led to the identification of the key enzymes involved in the catabolism of 2-fluorophenol (2-FP). A degradation pathway of 2-FP is proposed based on these enzymes and on the identification of metabolites with HPLC and 19F NMR. The initial step of 2-FP degradation involves hydroxylation of the aromatic ring to form 3-fluorocatechol by a two-component monooxygenase system of which one component provides reduced flavin adenine dinucleotide (FADH2) and the other catalyzes hydroxylation of 2-FP. Further degradation proceeds by ring cleavage with catechol 1,2-dioxygenase to form 2-fluoromuconate, which is then transformed by muconate cycloisomerase. A stoichiometric amount of fluoride was released and the products were further mineralized.

INTRODUCTION

The increased use of synthetic chemicals has led to the entry of toxic and recalcitrant compounds into the environment. Halophenols are prominent xenobiotics used in pharmaceutical, agricultural and other industrial applications (74). They are resistant to degradation and cause acute toxicity to various life forms (31). In general, fluorochemicals are gaining importance due to their stability, inertness and versatile possibilities for application in drugs and commercial products (3, 27, 29, 31, 51, 76).

During the last 30 years, degradation studies with haloaromatics have mainly been focused on chloro- and bromo-substituted compounds. In view of the concern over the presence of organofluorines in the environment, emphasis is now put also on the behavior and degradation of those compounds (31). The catabolism of only a few types of fluorinated compounds has been studied so far. Of the monofluoro aromatic compounds, the most investigated one is 4-fluorobenzoate, which can be used as a carbon source for growth by selected bacterial cultures (56, 66). Catabolism of fluorobenzene (13) and fluorophenols has been studied using pure strains of bacteria and fungi. Under aerobic conditions, Arthrobacter sp. strain IF1 transformed 4-fluorophenol into hydroquinone and released fluoride stoichiometrically (20, 21). Furthermore, 2-, 3-, and 4-fluorophenols were transformed to

fluorocatechols or fluoropyrogallols via hydroxylation catalyzed by phenol hydroxylase (6, 7, 8, 22, 37, 60).

Dehalogenation is the key step for decreasing the stability and inertness of a halogenated compound, which is accompanied by reducing toxicity and increasing biodegradability (52). In case of fluorophenols, fluoride release can occur before or after ring cleavage. Formation of the corresponding fluorocatechols and subsequent ring cleavage resulted in the release of > 99% of the fluoride in acclimated activated sludge during degradation of 2-, 3- and 4-fluorophenols (14, 22). Using pure strains of bacteria and fungi, release of 80% (8), 29% (60), 50% (37) and 9% (44) fluoride before ring cleavage has been reported when 2-fluorophenol was transformed into mixtures of catechol and fluorocatechol.

When 3-fluorocatechol is formed by hydroxylation of 2-FP, subsequent ring cleavage can result in the formation of fluoromuconic acids, which then may be degraded further via fluoromuconolactone to oxoadipate with release of fluoride (7, 71).

In this study, we describe the transformation of 2-fluorophenol by Rhodococcus sp.

strain FP1. We have analyzed the proteins involved in the degradation of 2-FP by MALDI-TOF/TOF mass spectrometry. We also have partially purified the enzymes involved in first two steps of 2-FP degradation and propose a degradation pathway.

MATERIALS AND METHODS

Chemicals. 2-Fluorophenol, catechol, 3-fluorocatechol, and 3-[(3-cholamidopropyl) dimethylammonio]-1-propanesulfonate (CHAPS) were purchased from Acros Organics (Belgium), and ampholytes was from GE Healthcare. To perform two-dimensional gel electrophoresis, immobilized pH gradient (IPG) dry strips were purchased from Amersham Biosciences. All chemicals were of analytical grade and used without further purification.

Analytical methods. Fluoride measurements were performed using a Dionex DX 120 ion chromatograph (Dionex, Sunnyvale, CA, USA) connected to an autosampler. The instrument was equipped with an Alltech A-2 anion column (100 × 4.6 mm, 7 µm) and an Alltech guard column (50 × 4 mm) operated at 30°C. An injection volume of 50 µl was used.

The eluent was a mixture of NaHCO3 and Na2CO3 in deionized water, with a flow rate of 0.8 ml min–1.

HPLC analyses were carried out using an Altima HP C18 reversed-phase column (100 mm × 2.1 mm, 3 µm particle size), in connection with Jasco PU-980 pumps, a Jasco MD-910

diode array detector and a Jasco UV-2075 detector. Samples of 10 µl were injected and compounds were isocratically eluted at a flow rate of 0.2 ml min-1. The mobile phase used was 70/30 (v/v) acetic acid/methanol containing 0.02 M ammonium acetate adjusted to pH 4.5 with glacial acetic acid. Spectra of eluted peaks were recorded between 200 and 600 nm.

19F NMR spectroscopy. The 19F Fourier transform spectra were recorded with a mercury plus console equipped with an Oxford/Varian 200 magnet and a Varian 4 nuc probe that was operated at a 19F observation frequency of 188.7 MHz, with 45° pulses and the inter scan delay set at 2 s. Typically 512 scans were recorded which yielded 16,212 data points with an acquisition time of less than 20 min.

Growth conditions. Cells of strain FP1 were grown aerobically at 30°C under rotary shaking conditions. Growth medium (MMY) contained per liter 5.37 g of Na2HPO4.12H2O, 1.36 g of KH2PO4, 0.5 g of (NH4)2SO4, 0.2 g of MgSO4.7H2O and 10 mg yeast extract (Difco laboratories). The medium was supplemented with a trace elements solution (5 ml l-1) that contained per l: 780 mg of Ca(NO3)2.4H2O, 200 mg of FeSO4.7H2O, 20 mg of Na2SeO4.10H2O, 10 mg of ZnSO4.7H2O, 10 mg of H3BO3, 10 mg of CoCl2.6H2O, 10 mg of CuSO4.5H2O, 4 mg of MnSO4.H2O, 3 mg of Na2MoO4.2H2O, 2 mg of NiCl2.6H2O and 2 mg of Na2WO4.2H2O.

Enrichment and isolation of a 2-FP degrading strain. The isolation of strain capable of growing on 2-FP was done by A. Duke as follows. Soil and rhizosphere samples collected from a contaminated site in Northern Portugal, which has received discharges of chemical industry effluents for more than 50 years (fine chemistry, agrochemicals), were combined as the initial inoculum for the 2-FP enrichments. This rhizosphere soil (approximately 5 g) was used to inoculate 250 ml flasks containing 50 ml of sterile MMY medium (11) and 2-FP was supplied to the liquid culture as sole carbon and energy source at a concentration of 50 mg l-1. Cultures were incubated on an orbital shaker (100 rpm) at 25°C.

Half of the suspension was removed and replaced with fresh medium at 6- to 7-day intervals.

Growth was monitored by measuring the optical density at 600 nm and liberation of fluoride was monitored using an ion-selective electrode. Bacterial strains were isolated from the enrichment culture by repetitive streaking onto Nutrient Broth (NB) agar medium. These pure cultures were then re-inoculated into MM containing 50 mg l-1 of 2-FP as the carbon source, and growth and fluoride release were monitored. A pure culture, designated strain FP1, was isolated and used for further study.

Identification of strain FP1. The 16S rRNA gene was amplified from genomic DNA by universal bacterial primers (Sigma-Genosys) 27F (5′-AGA GTT TGA TCM TGG CTC

AG-3′) and 1492R (5′-TAC GGY TAC CTT GTT ACG ACT T-3′) (39). The PCR mixture contained 10 µl of cell suspension (one colony of strain FP1 picked from an overnight LB agar plate and resuspended in 50 µl of water), 25 µl of Phusion PCR mix 2x (Finnzymes), and 0.25 µM forward and reverse primers in a final volume of 50 µl. PCR amplification was performed with a Techne thermocycler using the following conditions: initial denaturation for 11 min at 94°C, followed by 25 cycles of 98°C for 10 sec, 55°C for 30 sec, and 72°C for 15 sec, with a final extension step at 72°C for 5 min. The PCR products were verified by agarose (0.7%) gel electrophoresis, and purified with a Qiaquick PCR purification kit (Qiagen Inc.).

The purified PCR products were cloned into EcoRV digested pZero-2 vector (Invitrogen).

The products were transformed into electrocompetent E. coli Top10 cells and transformants were selected on LB plates containing kanamycin (50 µg ml-1) and X-gal for blue/white screening (47). The plasmids were extracted from 5 ml overnight cultures using High Pure Plasmid Isolation kit (Roche). The inserts were sequenced by GATC (Germany) and compared using BLAST (http://www.ncbi.nlm.nih.gov/BLAST/) for identification and phylogenetic classification.

Whole-cell transformation of 2-FP. Cells of strain FP1 were grown in MMY medium on 2-FP and harvested before the stationary phase was reached at an optical density of 450 nm of approximately 0.12. Following centrifugation at 4,000 × g for 8 min, cells were washed twice with 100 mM potassium phosphate buffer (pH 6.8) and resuspended with 5 ml of the same buffer. A cell suspension of strain FP1 was added to a 250 ml flask containing 100 ml of MMY medium supplemented with 1.15 mM 2-FP. Cells were incubated in a rotary shaker at 30°C and 200 rpm. Samples were taken with suitable time intervals, centrifuged at 16,000 × g, and supernatants were analyzed immediately by HPLC and ion chromatography.

Preparation of cell-free extract. Cells of strain FP1 were grown in MMY medium supplemented with 2-fluorophenol or succinate and harvested at mid-log phase by centrifugation (6,000 × g, 4°C for 30 min). The cells were washed twice with TEMG (50 mM Tris-SO4 + 1 mM EDTA + 1 mM β-mercaptoethanol + 5% glycerol, pH 7.0) and resuspended in the same buffer. Protease inhibitor cocktail (one tablet of Mini Complete per 10 ml suspension; Roche) was added to the cell suspension prior to sonication. The cells were subsequently broken using a Vibra-Cell sonicator (Sonics and Materials INC Danbury, Connecticut, USA). During sonication, the tubes were cooled on ice. To remove unbroken cells and debris, the preparation was centrifuged at 40,000 × g for 60 min. The cell-free extract thus obtained was either stored at -80°C or used immediately for enzyme assays. For

proteomic studies, cell-free extract was centrifuged at 80,000 × g for 20 min to remove membrane fractions. Protein concentrations were determined by using the Bradford method.

Two-dimensional gel electrophoresis (2-D). Two batches of cell-free extract were prepared by growing Rhodococcus sp. strain IF1 in separate flasks containing MMY medium supplemented with 2-FP. Controls were grown in flasks containing only succinate as a carbon source. To carry out proteomic analysis from cell-free extract, 200 µg of protein was incubated with nucleases at room temperature to remove nucleic acids. Protein samples were precipitated with four volumes of ice-cold acetone and subsequently incubated at -20°C for 1 h. Precipitated proteins were sedimented by centrifugation at 16,000 × g for 20 min at 4°C and resuspended in loading buffer (7 M urea, 2 M thiourea, 1% CHAPS (w/v), 0.5 % DTT (w/v), 0.5 % ampholytes (w/v)). The sample solution was applied on IPG strips of pH range 4-7 (Amersham, Uppsala, Sweden) by the overnight rehydration loading method. Strips were focused in an IPGphor instrument (GE Healthcare, Uppsala, Sweden) at 8,000 V for 6 h (48,000 Vh). The second dimension SDS-PAGE was run on a 12% polyacrylamide SDS gel with an Ettan DALT Twelve (GE Healthcare, Uppsala, Sweden). Protein spots were visualized by silver staining (Fig. 2A, 2B) (50) or Coomassie brilliant blue G-250 staining (Fig. 2C, 2D) (18).

For silver staining, gels were fixed with 50% methanol containing 12% acetic acid for 2 h, washed twice with 50% methanol for 30 min, and treated with 0.04% sodium thiosulfate for 1 min. Gels were washed with demineralized water and then impregnated in 0.1% silver nitrate solution for 20 min at 4ºC. After washing with demineralized water, gels were treated with 3% sodium carbonate solution containing 0.05% formaldehyde until the spots were visualized. The reaction was stopped with 1.4% EDTA solution and the stained gels were stored in 1% acetic acid.

Sample preparation for MALDI-MS and MS/MS analysis. Mass spectrometry experiments were done by H. Permentier. Selected spots from silver-stained and Coomassie-stained two-dimensional gel electrophoresis (2-D) were excised, deCoomassie-stained and digested with trypsin (Promega, Madison, WI, USA). For silver-stained spots, the destaining method was adopted from Kim et al. (34). For the Coomassie-stained gel, gel pieces were dried in Speed-Vac after washing twice with 25 mM ammonium bicarbonate and 50% acetonitrile. For tryptic digestion, dried gel pieces were swollen in 10 ng/µl trypsin solution and incubated at 37°C for 12 to 15 h. Peptides were recovered by adding a mixture of 75% acetonitrile and 25% of 5% formic acid in water.

Samples from digested proteins for MS were prepared by mixing 0.5 µl of the sample with 0.5 µl matrix solution (5 mg/ml α-cyano-4-hydroxycinnamic acid (CHCA) in 50%

acetonitrile containing 0.1% trifluoroacetate) and spotted on a stainless steel 192-well target plate. They were allowed to air dry at room temperature, and analyzed on a 4700 Proteomics Analyzer (Applied Biosystems, Foster City, CA, USA) MALDI-TOF/TOF mass spectrometer. For MS spectra, 1500 laser shots were acquired, and subsequently precursors from the resulting peptide spectra in the m/z range 840-4000 with a signal-to-noise threshold of 50 were automatically selected for analysis by MS/MS, with a maximum of 25 precursors per spot, excluding the most commonly observed peptide peaks of trypsin and keratin.

Protein identification. Peak lists of all MALDI-MS/MS spectra were produced by the 4000 Explorer software (version 3.5.3, Applied Biosystems) and submitted for search to Mascot (version 2.1, Matrix Science, London, UK) using the UniProt database restricted to Actinobacteria (www.uniprot.org, accessed on 15 June 2009). Search parameters were: 1 missed cleavage, methionine oxidation allowed as variable modification, precursor and fragment tolerance 150 ppm and 0.2 Da, respectively. Proteins are reported if they have a significant score (p < 0.05) and are identified with at least two peptides.

Protein purification. Cell-free extract of strain FP1 induced with 2-FP was loaded onto a DEAE Sepharose column (50 ml; Pharmacia, Uppsala, Sweden) pre-equilibrated with TEMG buffer (50 mM Tris-SO4, 5% glycerol, 1 mM β-mercaptoethanol and 0.5 mM EDTA, pH 7.0). After washing the column with two column volumes of buffer, elution was carried with a linear gradient from 0 - 100% of 0.5 M (NH4)2SO4 in the same buffer. Active fractions were collected, separately concentrated by using 10 kDa Millipore filters (Amicon, USA),

and tested for monooxygenase and dioxygenase activities. Fractions containing 2-fluorophenol monooxygenase and catechol 1,2-dioxygenase activities were desalted and

loaded separately onto a gel filtration column (Superdex 200, 24 ml bed volume). Proteins were eluted with a buffer (pH 7.0) containing 100 mM NaCl, 1mM EDTA and 1 mM β-mercaptoethanol at a flow rate of 0.2 ml/min. Fractions were assayed for activity. To check the purity, concentrated fractions were run on 12% sodium dodecyl sulfate (SDS)-polyacrylamide gel and stained with Coomassie Brilliant Blue R-250.

Enzyme assays. Monooxygenase activity with 2-FP was measured in 50 mM Tris-Cl buffer (pH 7.5). Reaction mixtures contained 1 mM ascorbate, 5 µM FAD, 3 µg of flavin reductase (FpdB, purified previously (21)), 180 U ml-1 catalase (Fluka, from bovine liver), 1.2 mM NADH and 600 µM of 2-FP. Time course conversions were carried out in 2000 µl

and were started by the addition of NADH. Samples of 200 µl were taken with time intervals of 10–25 min and quenched by addition of an equal volume of HPLC eluent (described above). Supernatants of vortexed and centrifuged samples were analyzed with HPLC.

Catechol 1,2-dioxygenase activities were measured using a Perkin-Elmer Lambda Bio 40 spectrophotometer at 25ºC by monitoring the formation of corresponding cis,cis-muconate at 260 nm (εcis,cis-muconate = 16,800 M−1 cm−1, ε2-fluoro-cis,cis-muconate = 14,900 M−1 cm−1, ε

3-fluoro-cis,cis-muconate = 14,900 M−1 cm−1) (16) in an assay mixture containing 50 mM Tris-Cl buffer (pH 7.5), 400 µM catechol or substituted catechol, and the appropriate amount of enzyme.

RESULTS

Isolation and characterization of cultures capable to degrade 2-fluorophenol. A pure bacterium (strain FP1) capable of aerobic biodegradation of 2-FP was isolated after 4 months of selective enrichments. Strain FP1 was able to use the compound as the sole carbon and energy source. Biodegradation was detected by growth and stoichiometric fluoride release. When plated on NA, small white colonies appeared and when analysed under a light microscope gram-positive, rod shaped cells were observed.

Strain FP1 was identified by 16S rRNA gene analysis as belonging to the genus Rhodococcus. According to BLAST results and subsequent phylogenetic analysis, it became evident that the rRNA sequence of FP1 clusters with that of Rhodococcus sp. TCH4 (AB183439) and Rhodococcus sp. TCH14 (AB183440), which were isolated from trichloroethylene-contaminated soil and are able to degrade o-xylene (73). The sequence also clustered with the corresponding 16S rRNA gene sequence of Rhodococcus opacus strain ML0004 (DQ474758), which was isolated from soil and studied because it produces epoxide hydrolase (40).

Degradation of 2-FP by strain FP1. Rhodococcus sp. strain FP1 completely degraded 1.15 mM 2-FP in MMY medium in 110 h. The optical density increased up to OD450 0.14 and stoichiometric release of fluoride was observed. The highest specific growth rate, which occurred in the early mid growth phase, was 0.012 h-1 (Fig. 1). During degradation of 2-FP, transient accumulation of 3-fluorocatechol (3-FC) was observed by HPLC. Further increase in optical density at 450 nm up to 0.15 in 70 h suggested formation of one or more non-fluorinated intermediates during degradation of 2-FP which were utilized after 2-FP was depleted. Control flasks containing the same amount of 2-FP were incubated

in parallel. A drop in 2-FP concentration of 2-6% due to volatilization, absorption or chemical decomposition in control flasks was measured.

Identification of proteins induced with 2-FP in strain FP1 by MALDI-MS/MS.

To explore the degradation pathway of 2-FP, we looked for the differential induction of proteins of Rhodococcus sp. stain FP1 grown with 2-FP compared to succinate-induced proteins. Cell-free extract from strain FP1 grown on 2-FP was analyzed by running two-dimensional gels and extract from succinate-grown cells was used as a control.

0 50 100 150 200 present at a higher level than in extracts prepared from succinate-grown cells (Fig. 2). From the 2-D gel of 2-FP-induced proteins, 120 spots from a silver-stained gel and 109 spots from a Coomassie-stained gel were selected, excised, destained and digested with trypsin. For 98 spots in silver-stained gel and 70 spots in the Coomassie-stained gel, we observed good spectra (<900 Da peptide mass), with peak intensities suitable for MALDI-MS/MS analysis.

The rest of the spots could not be used for MS/MS due to low intensities of the peptide peaks.

The fragmentation spectra obtained from the prominent spots were used for protein identification by database searching. The Actinobacteria protein database was used to identify the induced proteins because strain FP1 was a Rhodococcus strain. Although the genome of strain FP1 has not been sequenced and the encoded proteins are therefore not represented in the Actinobacteria protein database, good matches were obtained with proteins from Rhodococcus strain RHA1 and Rhodococcus opacus strain B4, indicating the presence of homologous genes in these organisms.

FIG. 1. Growth of strain FP1 in MMY in a 1 l flask supplemented with 1.14 mM 2-FP. Symbols: ■, 2-FP concentration; ▼, 3-FC concentration;

●, optical density at 450 nm; ▲, F concentration.

FIG. 2. Two-dimensional gel electrophoresis of silver-stained protein pattern (A and B) and Coomassie-stained protein pattern (C and D) from Rhodococcus sp. strain FP1 grown with the addition of succinate (A and C) and 2-FP (B and D) in MMY medium. Selected protein spots with increased levels are highlighted by numbered arrows. They correspond to the proteins listed in Table 1 (from Coomassie-stained gel) and Table 2 (from silver-stained gel).

Of the identified proteins, phenol 2-monooxygenase, phenol hydroxylase, catechol 1,2-dioxygenase and muconate cycloisomerase are directly associated with the degradation of 2-FP. This clearly indicates that strain FP1 selectively induced ortho-cleavage enzymes when exposed to 2-FP. In the 2-D gel, proteins of spot 1, 2 and 3 (Fig. 2D) of ~60 kDa were similar to phenol 2-monooxygenase of Rhodococcus sp. strain RHA1 (Q0SE51), while the protein of spot 4 was matched to a putative phenol hydroxylase of Rhodococcus opacus B4 (C1B264).

Formation of this triplet of spots with the same molecular weights and slightly different pI values may be caused by either posttranslational modification or minor amino acid differences (57).

TABLE 1. Proteins with increased levels in Rhodococcus sp. strain FP1 grown in MMY medium supplemented with 1 mM 2-fluorophenol as compared to 1 mM succinate.

Spot

no.a Protein name Identified peptides Mascot score b

Mr

(kDa), pI

Accession no./strain 1 Phenol 2-monooxygenase VEDVTTHPAFR

VLYWNHAIINPPVDR

2 Phenol 2-monooxygenase VEDVTTHPAFR VLYWNHAIINPPVDR

3 Phenol 2-monooxygenase VEDVTTHPAFR

SSFTQNAAVMGTPFDYPLSSR

4 Putative phenol hydroxylase VKDVTTHPAFR

TSYTQQAAVMGSPFDYPLSSR

5 Catechol 1,2-dioxygenase HGVTYPEYR

GSIEGPYYIENSPELPSK GGEWIDSDVASATKPELILDPK

176 31, 4.7 Q0SE58/Rhodococcus sp. RHA1

6 Catechol 1,2-dioxygenase HGVTYPEYR

GSIEGPYYIENSPELPSK

140 31, 4.7 Q0SE58/Rhodococcus sp. RHA1

7 Catechol 1,2-dioxygenase HGVTYPEYR

GSIEGPYYIENSPELPSK

9 Superoxide dismutase SVYTLPELPYDYAALEPHISGK 99 23, 5.3 C1B8Y4/Rhodococcus opacus B4

a Spot number according to Fig. 2D.

b Mascot score represents the probability that the observed match is a random event. Only protein scores with P values of < 0.05 are reported.

The sequences of Q0SE51 and C1B264 are very homologous (with 92% similarity, sequences obtained from UniProtKB/TrEMBL), but we have identified 7 different peptides which clearly distinguish the two proteins (Fig. 3). Phenol 2-monoxygenase and the putative phenol hydroxylase are annotated as different enzymes from different species of

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