The oxygenase component of phenol hydroxylase from Acinetobacter radioresistens S13 Sara Divari 1 , Francesca Valetti 1 , Patrizia Caposio 4 , Enrica Pessione 1 , Maria Cavaletto 2 , Ersilia Griva 1 , Giorgio Gribaudo 4 , Gianfranco Gilardi 1,3 and Carlo Giunta 1 1 Dipartimento di Biologia Animale e dell’Uomo, Universita ` di Torino, Italy; 2 Dipartimento di Scienze e Tecnologie Avanzate, Universita ` del Piemonte Orientale, Alessandria, Italy; 3 Department of Biological Sciences, Imperial College of Science, Technology and Medicine, London, UK; 4 Dipartimento di Sanita ` Pubblica e Microbiologia, Universita ` di Torino, Italy Phenol hydroxylase (PH) from Acinetobacter radioresis- tens S13 represents an example of multicomponent aromatic ring monooxygenase made up of three moieties: a reductase (PHR), an oxygenase (PHO) and a regulative component (PHI). The function of the oxygenase component (PHO), here characterized for the first time, is to bind molecular oxygen and catalyse the mono-hydroxylation of substrates (phenol, and with less efficiency, chloro- and methyl-phenol and naphthol). PHO was purified from extracts of A. radio- resistens S13 cells and shown to be a dimer of 206 kDa. Each monomer is composed by three subunits: a (54 kDa), b (38 kDa) and c (11 kDa). The gene encoding PHO a(named mopN) was cloned and sequenced and the corresponding amino acid sequence matched with that of functionally related oxygenases. By structural alignment with the cata- lytic subunits of methane monooxygenase (MMO) and alkene monooxygenase, we propose that PHO a contains the enzyme active site, harbouring a dinuclear iron centre Fe-O-Fe, as also suggested by spectral analysis. Conserved hydrophobic amino acids known to define the substrate recognition pocket, are also present in the a-subunit. The prevalence of a-helices (99.6%) as studied by CD confirmed the hypothized structural homologies between PHO and MMO. Three parameters (optimum ionic strength, tem- perature and pH) that affect kinetics of the overall phenol hydroxylase reaction were further analyzed with a fixed optimal PHR/PHI/PHO ratio of 2/1/1. The highest level of activity was evaluated between 0.075 and 0.1 M of ionic strength, the temperature dependence showed a maximum of activity at 24 °C and finally the pH for optimal activity wasdeterminedtobe7.5. Keywords: multicomponent monooxygenase; phenol hydroxylase; purification; molecular cloning; catalytic subunit. Phenol-degrading aerobic bacteria are able to convert phenol into nontoxic intermediates of the tricarboxylic acid cycle via an ortho or meta pathway [1]. The first step of both routes is the monohydroxylation of the ortho position of the aromatic ring [2]. The enzyme responsible for this reaction is the monooxygenase phenol hydroxylase (PH). Aromatic monooxygenases are divided into two groups: activated-ring monooxygenases (monocomponent) and nonactivated-ring enzymes (multicomponent). In the latter case, the active site must contain a strong hydroxyl- generating-unit, i.e. a dinuclear iron centre in which an oxygen atom is complexed with two iron atoms Fe-O-Fe (while in the former case the enzyme is a simple flavoprotein [3,4]). Furthermore, a short redox chain is required to supply electrons from NAD(P)H to the dinuclear iron centre itself. Such a multicomponent organization is present in a number of enzymes that are able to hydroxylate and start the detoxification process of poorly reactive aromatics and aliphatics, often recalcitrant to degradation. Among these molecules examples are toluene, that is converted to p-hydroxytoluene by toluene-4-mono-oxygenase in Pseudo- monas mendocina KR1 [5]; xylene, the substrate of a xylene/ toluene monooxygenase in Pseudomonas stutzeri OX1 [6]; methane, that is converted to methanol by methane monooxygenases in Methylococcus capsulatus Bath [7], Correspondence to C. Giunta, Dipartimento di Biologia Animale e dell’Uomo, Universita ` di Torino, Via Accademia Albertina, 13, 10123 Torino, Italy. Fax.: +39 0116704692, Tel.: +39 0116704644, E-mail: carlo.giunta@unito.it Abbreviations: AMO, alkene monooxygenase; AMOa, alkene mono- oxygenase a subunit from Nocardia corallina B-276; AMO Py2, alkene monooxygenase from Xanthobacter Py2; C1,2O, catechol 1,2 di- oxygenase; MMO M, methane monooxygenase; MMO B, methane monooxygenase from Methylococcus capsulatus Bath; MMOMz, M methane monooxygenase from Methylosinus trichosporium OB3b; PH, phenol hydroxylase; PHI, phenol hydroxylase regulatory protein; PHR, phenol hydroxylase reductase; PHO, phenol hydroxylase oxygenase; T3MO, toluene-3-monooxygenase from Pseudomonas pickettii PKO1; T4MO, toluene-4-monooxygenase from Pseudomonas mendocina KR1; Xyl/TMO, xylene/toluene monooxygenase from Pseudomonas stutzeri. Enzymes: alkene monooxygenase (EC 1.14.13 ); phenol hydroxylase (EC 1.14.13.7); toluene 4-monooxygenase (EC 1.14.14.1); toluene 3-monooxygenase (EC 1.14.13 ), xylene/toluene monooxygenase (EC 1.14.14.1); methane monooxygenase (EC 1.14.13.25). (Received 18 December 2002, revised 19 March 2003, accepted 26 March 2003) Eur. J. Biochem. 270, 2244–2253 (2003) Ó FEBS 2003 doi:10.1046/j.1432-1033.2003.03592.x Methylosinus thrichosporium [8] and Methylocystis [9]; alkenes, converted to the corresponding epoxides by the alkenes monooxigenase of Nocardia corallina B-276 [10]. Surprisingly, in a few bacterial strains [11,12], a very similar system was also described to recognize phenol, even if in this case the aromatic ring to be processed is activated by the hydroxyl group and a simple FAD-dependent monocom- ponent enzyme would be able to catalyze the reaction, as reported for Pseudomonas pickettii [13] and Bacillus stearo- thermophylus [14]. The multimeric phenol hydroxylase from Pseudomonas sp. strain CF 600 [1,15,16] is of particular interest because its biochemical and genetic characteri- zation indicated an organization very similar to both methane monooxygenase from methanotrophs [17–20] and to alkene monooxygenase from Nocardia corallina [21]. All these monooxygenases are composed of one NADH binding monomeric reductase, one multimeric (abc) 2 oxygenase containing the dinuclear iron-centre (on the a subunit) and binding oxygen and substrate, plus a small regulatory component whose NMR structure has been reported recently [22–24]. Among the Acinetobacter genus, only genetic data are available for the phenol- degrading A. calcoaceticus NCIB 8250 [12]. These show a genomic organization of the PH encoding genes very similar to that reported for P. sp. strain CF600 in addition to a significant amino acid sequence homology. Therefore there are at least two known PHs expressed by different bacterial genera with a multicomponent organization. In a previous paper, we showed that PH from Acineto- bacter radioresistens LMG S-13648 (thereafter, designed to as A. radioresistens S13) consists of three components [25]. The reductase component, PHR, is a monomeric iron- sulfur-flavoprotein [25] that transfers reducing equivalents from NAD(P)H to the oxygenating moiety PHO described in this work, and PHI is an intermediate component necessary for catalysis [26]. This multicomponent enzyme confers to the organism a very high (100 mgÆL )1 Æ h )1 ) phenol degradation rate when this substrate is the sole carbon and energy source. The potential of this organism in bioremediation is shown by its ability to grow on activated sludges of industrial wastes, from which the strain was isolated by our group [27]. This paper describes the purification, the characterization and the catalytic properties of the oxygenase component of PH from A. radioresistens S13, as well as the molecular cloning of the a-subunit responsible for catalysis. Materials and methods Cell growth and preparation of soluble extract A. radioresistens S13 cells were grown in a Sokol and Howell [28] minimal medium in which phenol was the sole carbon source. Phenol was added in a fed-batch fermenta- tion procedure (100 mgÆL )1 Æh )1 ) and the culture was incubated at 30 °C for 23–24 h. Cells were harvested by ultracentrifugation at 15 000 g,washedtwicein50mL Hepes/NaOH buffer, pH 7.0 and stored frozen ()80 °C). Biomass (200 g per 200 mL) in 50 m M Hepes/NaOH buffer, pH 7.0 were sonicated on ice for a total time of 20 min at 20 kHz with intervals of 1 min (Microsonix Sonicator Ultrasonic Liquid Processor XL2020). The obtained soluble extracts were then centrifuged at 100 000 g for 1 h at 4 °C (ultracentrifuge LB60M Beckman). The supernatants were considered as the crude extracts. Phenol hydroxylase assay PH activity was measured polarographically by means of a Clark-type electrode (YSI Model 5300). The assay was carried out in the presence of 1.68 m M NADH, 0.6 l M PHO, 1.2 l M PHR, 0.6 l M PHI and 100 m M Mops/NaOH buffer, pH 7.4 at 24 °C. The reaction was started by adding 1m M phenol (Fluka). The same experiment was performed using the following substrates (1 m M ): p-cresol, m-cresol, 3,4-dimethylphenol, b-naphthol, a-naphthol, 3-chloro- phenol, 4-chlorophenol, 3,4-dihydroxyphenol, p-hydroxy- benzoic acid, m-hydroxybenzoic acid, 2,4-dinitrophenol, 2,4-dichlorophenol, 3,4-dichlorophenol, 2,4,5-trichloro- phenol, 2,2¢-dihydroxybiphenyl and L -tyrosine. Poorly water soluble compounds were prepared as 300-fold con- centrated stock solutions in methanol and only 10 lLwere added to the reaction mixture. No interference or protein damage was observed due to the presence of this amount of methanol. Kinetics of phenol consumption by PH reconsti- tuted complex were also evaluated by a discontinuous assay at 24 °C, in 100 m M Mops/NaOH buffer pH 7.4 at the same reactant concentration used for the oxygen consump- tion assay (1.68 m M NADH, 0.6 l M PHO, 1.2 l M PHR, 0.6 l M PHI and 1 m M phenol). Residual phenol concen- tration at different times of reaction was evaluated by HPLC (Merck Hitachi L6200 system) on a Lichrosphere RP-18 100 column (Merck) equipped with a precolumn (Lichrosphere RP-18 40 Merck). An isocratic separation (50% acetonitrile) at a flow rate of 1 mLÆmin )1 was performed in order to separate the phenol peak that was monitored at 270 nm by a Diode Array detector system (Merck Hitachi L4500) and quantitated by the correcting peak area against the total area. The obtained data were fitted to a biexponential decay function and only the faster component was considered for turnover number calcula- tion. The same procedure was applied to the other tested substrates, adjusting HPLC eluents to pH ¼ 3.0 with concentrated H 2 SO 4 , and varying the monitored wave- length to the corresponding k-max. A third kind of assay was based on the continuous measurement of NADH consumption. The same reaction mixture described above was monitored at 24 °C for NADH disappearance at 340 nm with a Beckman DU 70 spectrophotometer and the kinetics calculated by taking the initial velocity as the tangent to the obtained curve. The reaction was started by substrate addition and corrected for basal NADH decay. Purification of PHO The crude extract was applied on an anion exchange DE52- cellulose (Whatman) column (2.6 · 20 cm) equilibrated previously with 50 m M Hepes/NaOH buffer pH 7.0. The elution was obtained with a 0–0.5 M sodium sulphate gradient in 50 m M Hepes/NaOH buffer pH 7.0 (final volume, 1.1 L). After ultrafiltration (membrane Diaflo, cut off 30 kDa; Amicon), the PHO containing fractions were applied on a Q-Sepharose FF (Pharmacia) column (1.3 · 26 cm) equilibrated with 50 m M Hepes/NaOH Ó FEBS 2003 A. radioresistens S13 phenol hydroxylase (Eur. J. Biochem. 270) 2245 buffer, pH 7.0 containing 0.15 M sodium chloride. PHO was eluted from this column with a 0.15–0.35 M NaCl linear gradient (total volume 1.2 L) in 50 m M Hepes/NaOH buffer pH 7.0. Active fractions eluted were concentrated by ultrafiltration and applied on a hydrophobic Phenyl- Sepharose 6FF column (Pharmacia) (1.3 · 13 cm). The column was equilibrated with 50 m M Hepes/NaOH buffer pH 7.0, containing 0.15 M sodium sulphate and the sample adjusted to the corresponding ionic strength with the same buffer. A mixed gradient was applied with an initial linear section from 0.15–0 M Na 2 SO 4 in 50 m M Hepes/NaOH buffer pH 7.0 (50 mL), an isocratic step of 120 mL of 50 m M Hepes/NaOH buffer pH 7.0, a 50-mL linear gradi- ent from 50 m M Hepes/NaOH buffer pH 7.0 to water and a final isocratic step in 100% water. PHO was eluted in this last section of the mixed gradient. In order to avoid prolonged exposure of PHO to water, the 3 mL eluted fractions were collected in tubes containing the same volume of 200 m M Hepes/NaOH buffer pH 7.0. PHO fractions in 100 m M Hepes/NaOH buffer, pH 7.0 were concentrated, as described previously, and stored at )80 °C until required. An average yield of 0.1 lmol of purified protein was obtained from 200 g of biomass following such procedure. Characterization Total protein concentration was estimated colorimetrically by the Bradford method [29] using BSA as standard. Molecular mass was determined by gel filtration and SDS/PAGE. A Superdex 200-FPLC (Pharmacia) column (1.6 · 60 cm) was equilibrated with 50 m M Hepes/NaOH buffer pH 7.0 with 0.05 M Na 2 SO 4 and calibrated with thyroglobulin (669 kDa), ferrytin (440 kDa), catalase (232 kDa), aldolase (158 kDa), BSA (67 kDa), ovalbumin (43 kDa) and chymotrypsinogen A (25 kDa) as standards. The molecular mass of PHO subunits was also determined by means of SDS/PAGE, performed on 12.5% poliacryl- amide gel. The molecular mass standards were: phosphory- lase b (97 kDa), BSA (67 kDa), ovalbumin (45 kDa) carbonic anhydrase (31 kDa), trypsin inhibitor (21 kDa) and lysozyme (14 kDa). The separation of the protomers was achieved by RP-HPLC. PHO (32 l M )wasmixedto 178 lL of 50% acetonitrile solution and 1% formic acid. Protomers generated were purified using a HPLC Merck- Hitachi L6200 system equipped with a Lichosphere 100 RP-8 column (Merck). The flow rate was 1 mLÆmin )1 .The column was equilibrated with solvent A (water/trifluoro- acetic acid, 100/0.08) and protomers were eluted using a linear gradient of 20–100% solvent B (water/acetonitrile/ trifluoroacetic acid, 10/90/0.08) over 80 min. N-Terminal sequences of the three subunits of PHO (a,b,c) were deter- mined after SDS/PAGE and Western Blotting onto Immo- bilon P membrane. The protein bands were sequenced following the Edman degradation, using a 470-A phase sequencer (Applied Biosystems USA). The isoelectric point was determined by analytical IEF electrophoresis (Phast- System, Pharmacia); the markers were those supplied by Pharmacia (pI calibration kit). The iron and sulfur content was determined by colorimetric methods as described previously [30,31]. CD measurements were performed on a Jasco Spectropolarimeter J-715 equipped with tempera- ture-controlled Peltier Jasco PTC-348WI. Spectra were acquired in the region 190–260 nm with a 0.1-cm path length cell and a scan speed of 20 nmÆmin )1 with a response time of 8 s. The CD spectrum reported is the average of three scans at 0.5 nm resolution and a bandwidth of 2 nm. The concentration was 1.25 l M for PHO. The spectrum was analyzed with the CDNN deconvolution program [32]. Construction and screening of the genomic library The A. radioresistens S13 genomic library was prepared as described [33]. Briefly, purified genomic DNA (10 lg) was partially digested with Sau3AI, and DNA fragments (10–20 kb) were fractionated by preparative gel electro- phoresis. Thereafter, the partially digested genomic DNA was partly completed with the Klenow fragment of E. coli DNA polymerase I and dGTP and dATP, and then ligated to LambdaGEM-12 XhoI half-site arms (Promega) accord- ing to the manufacturer’s instructions. The A. radioresistens S13 genomic library resulted in 1.3 · 10 5 independent recombinant clones that were stored and screened with- out amplification. It was plated on layers of susceptible bacteria host at a density of 5000 plaques per 150-mm diameter Petri dish, and a duplicate set of nylon filters (Hybond-N + , Amersham) was taken from each filter. Filters were hybridized with a radiolabeled DNA probe in a solution containing 5 · NaCl/Cit-5 · Denhardt’s-1% SDS-100 lgÆmL )1 of denatured salmon sperm DNA for 18 h at 65 °C. Filters were then washed twice for 20 min at 65 °Cin2· NaCl/0.1% Cit SDS and then twice for 20 min at 65 °Cin0.2· NaCl/Cit-0.1% SDS. DNA inserts from recombinant phages of interest were prepared, restricted with EcoRI and subcloned into pGEM-7Zf(+) (Promega). DNA templates were then sequenced by the dideoxy-chain terminator method using a Sequenase 2.0 DNA sequencing kit (USB). Sequence data were used for homology search in the GenBank database and have been deposited in the same database with accession number for AF521658. The hybridization probe was prepared by means of PCR using degenerated primers synthesized according to the amino acid sequences, SQVKTTVKKL and LLSIVAMMM. PCR was carried out with 100 ng of A. radioresistens S13 genomic DNA in PCR buffer (10 m M Tris/HCl, pH 8.3, 50 m M KCl, 1.5 m M MgCl 2 ) containing the two primers (0.250 l M each), the four dNTPs (200 l M each) and 2.5 U of Taq DNA polymerase (Perkin-Elmer). Samples were subjected to 30 cycles of amplification with the following cycle profile: denaturation at 94 °Cfor1min, annealing to 50 °C for 1 min, extension at 72 °Cfor2min. pH, temperature and ionic strength dependence of enzyme activity PHO activity was determined at 24 °C in the pH range 5.5–9, using Good’s buffers (50 m M Mes-Mops-CHES) adjusted at a fixed ionic strength of 0.119 M by addition, where necessary, of NaCl. No difference was observed in the reaction upon adjusting ionic strength by buffer concentra- tion or by addition of NaCl. The activity was evaluated as oxygen consumption by a Clark electrode. The temperature dependence of phenol hydroxylation reaction at pH 7.4 was investigated in the range 20–37 °C by means of a thermo- stated reaction cell. Gay–Lussac correction factors were 2246 S. Divari et al. (Eur. J. Biochem. 270) Ó FEBS 2003 employed for calculating the saturating oxygen concentra- tion, at the tested temperature, in the aqueous reaction mixture. Ionic strength dependence of PH activity was studied in the same reaction condition described above, at 24 °C and pH 7.4. The ionic strength was varied from 4–164 m M by adjusting the concentration of Mops/NaOH buffer in a range 10–250 m M . The experiment was repeated in Hepes/NaOH buffer pH 7.4 in the same concentration range. The precise ionic strength at each buffer concentra- tion was calculated by means of the on-line program available at http://www.bi.umist.ac.uk/users/mjfrbn/buffers/ makebuf.asp [34]. Data obtained from the two buffer systems were found coincident within the experimental error and therefore gathered in the same dataset. Perfectly comparable results were also obtained when the ionic strength was adjusted to the desired value by NaCl addition. Results PHO purification Purification of PHO was achieved by two anion exchange columns followed by a hydrophobic interaction chroma- tography. The molecular mass value of the purified PHO was estimated to be 207 kDa by gel-permeation, whilst three bands of 54 kDa, 37.8 kDa and 11.6 kDa were observed on SDS/PAGE gels (Fig. 1A). This suggests a hexameric structure such as (abc) 2. The three subunits were also separated by RP-HPLC with a weaker interaction of c subunit to the C8 column matrix and a higher affinity of the b and a subunits (Fig. 1B). The isoelectric point of the purified PHO was determined to be 6.74. N-terminal primary structure alignment N-terminal sequences of the three subunits of PHO were determined to be SQVKTTVKKL for the a-, TLEIKTA GIE for the b- and SVRAIRPDYD for the c-subunit. These sequences were compared with analogous sequences of PHOs extracted from different bacterial species that express multicomponent phenol hydroxylases. The degree of identity is very high (60%) for the a-andb-subunits of A. radioresistens S13 and A. calcoaceticus [12], lower (40–20%) between A. radioresistens S13 and Pseudomonas sp. strain CF600 [11] and very low (30–20%) among the three considered c-subunits (A. radioresistens S13, A. calcoaceticus and P. sp. strain CF600). Fig. 1. The multimeric structure of PHO from A. radioresistens S13. (A) SDS/PAGE of PHO from different purification steps. Lane 1, molecular mass standards; lane 2, crude extract; lane 3, after DE52-cellulose; lane 4, after Q-Sepharose; lane 5, after phenyl sepha- rose. The molecular masses of the protomers a, b, c are54kDa,38kDaand11kDa, respectively. (B) PHO protomer separation by RP-HPLC. PHO (32 l M ) was mixed to 178 lLof50%H 2 O-50% acetonitrile solution and 1% formic acid. Protomers generated were purified using a HPLC Merk-Hitachi L6200 system equipped with a Lichosphere 100 RP-8 column (Merk). The flow rate was 1mLÆmin )1 . The column was equilibrated with solvent A (H 2 O/Trifluoroacetic acid, 100/0.08) and protomers were eluted using a linear gradient of 20–100% solvent B (H 2 O/ CH 3 CN/trifluoroacetic acid, 10/90/0.08) over 80 min. Ó FEBS 2003 A. radioresistens S13 phenol hydroxylase (Eur. J. Biochem. 270) 2247 Cloning and nucleotide sequence of the phenol hydroxylase PHO a-subunit The gene encoding for the PHO a-subunit was obtained from a genomic library of A. radioresistens S13, prepared and screened with a specific DNA probe obtained by PCR amplification of genomic DNA with degenerated oligo- nucleotides designed on the NH 2 -terminal sequence and on peptides derived by trypsin digestion of the purified protein. After subcloning and sequence analysis, the nucleotide and deduced amino acid sequences were matched with protein sequences. To complete the cloning of the gene of the PHO a-subunit, about 10 4 phage plaques were screened with the PCR product as probe and the P19.2 clone was selected for PHO a-encoding sequences. The gene encoding for the PHO a-subunit (mopN)waslocatedona1.9-kbEcoRI segment of the P19.2 clone and the nucleotide sequence was determined. The sequence of the 1.9 kb EcoRI fragment revealed that the mopN gene of A. radioresistens S13 encodes an ORF of 1527 base pairs corresponding to a polypeptide of 509 amino acids with a calculated molecular weight of 59.9 kDa. This value is in agreement (within the experimental error) with that obtained by SDS/PAGE resolution of purified PHO subunits. Further analysis (Table 1) of the whole deduced amino acid sequences revealed a high degree of similarity to other bacterial oxo- iron oxygenase so far characterized [6,17,18,35–38], inclu- ding the typical EXXH motif involved in the coordination of the catalytic dinuclear iron centre, as already described in MMO from M. capsulatus and M. trichosporium and in PHOs from Pseudomonas CF 600 and A. calcoaceticus NCIB8250 (Table 1). Furthermore, many hydrophobic residues of the protein aligned with conserved amino acids of MMO active site pocket (Table 1). PHO a shows the highest degree of identity (92%) with the DMS oxygenase component of A. sp. strain 20B (accession number BAA2333.1) [39] and with the PHO component 4 (ORF 4) of A. calcoaceticus NCBI 8250 (accession number CAA85383.1) [12]. High similarity (70% identity) was also observed with both the PHO component D of P. putida strain H (accession number CAA56743.1) [40], and the P3 protein of the multicomponent PHO of Pseudomonas sp. strain CF600 (accession number AAA5942.1) [11]. More- over, A. radioresistens S13 PHO a showed 68% and 65% identity with a PHO component of Ralstonia sp. K1 (accession number BAA84121.1) [41] and with the toluene ortho-monooxygenase (tomA) of Burkholderia cepacia G4 (accession number AAK07411.1). Spectroscopy analysis of purified PHO The UV/vis spectrum of purified PHO shows a single peak at 280 nm (Fig. 2). The e 280 nm wascalculatedtobe 643 800 M )1 Æcm )1 in 50 m M Hepes/NaOH buffer, pH 7.0. In the visible region, a broad shoulder is highlighted when comparing holo- and apo-PHO (Fig. 2, inset). This can be attributed to the presence of oxo-bridged di-iron centres as reported recently [16] for phenol hydroxylase from P. sp. strain CF600. For the latter, the extinction coefficient at 350 is between 4800 and 6000 M )1 Æcm )1 per di-iron centre. Using these values, we can estimate the presence of 1.9– 2.4 mol of di-iron centre per mol of dimer (abc) 2 ,closeto Table 1. Sequence alignment of A. radioresistens S13 PHO a with the active site and metal coordinating regions of homologous di-nuclear oxo-iron oxygenases. Conserved residues involved in the oxo-iron centre coordination are indicated in bold. Hydrophobic residues delimiting the active site cavity are underlined. Residues are numbered only for the enzymes with known 3D structures [19,20]. PHO a, A. radioresistens S 13 phenol hydroxylase a component; MMO B, M. capsulatus methane monooxygenase [17]; MMO M, M. trichosporium methane monooxygenase [18]; AMO a, N. corallina alkene monoxygenase; AMO Py2, X. Py2 alkene monoxygenase [36]; Xyl/TMO, P. stutzeri xylene/toluene monooxygenase [6]; T4MO, P. mendocina toluene 4 monooxygenase [37]; T3MO, P. pickettii toluene 3 monooxygenase [38]. Enzyme Residue Sequence Residue Sequence Residue Sequence Residue Sequence Residue PHO a – LEYQAFQG –/– MQSIDELRHV –/– FEFLLAISFAFEYVLTNLLFV –/– TFGFSAQSDEARHMTLG || ::: | :||: | | : | | : : :: | :: || |:| |::: | :::||: | || | Mmo B 110 LEVGEYNA 117/139 AQVLDEIRHT 148/198 VECSLNLQLVGEAGFTNPLIV 218/234 TVFLSIETDELRHMANG 250 Mmo M 110 GEYNAIAA 117/139 AQVLDEIRHT 148/198 VECSVNLQLVGDTCFTNPLIV 218/234 TVFLSVRTDELRHMANG 250 Amo a – LTNAEYQA –/– AQMLDEVRHA –/– LDVIIDLNIVAETAFTNILLV –/– SVFLSIQSDEARHMANG Amo Py2 – VEHMAVTM –/– FGMLDETRHT –/– VEAALATSLTLEHGFTNIQFV –/– NLLSSIQTDEARHAQLG Xyl/TMO – EEYAASTA –/– FGMMDENRHG –/– VAVSIMLTFAFETGFTNMQFL –/– SLISSIQTDESRHAQQG T4MO – GEYAAVTG –/– FGMMDELRHG –/– ISVAIMLTFSFETGFTNMQFL –/– NLISSIQTDESRHAQQG T3MO – GEYAAMSA –/– FGMLDENRHG –/– IDIAIMLTFAFETGFTNMQFL –/– SLISSIQTDESRHAQIG 2248 S. Divari et al. (Eur. J. Biochem. 270) Ó FEBS 2003 the expected value of one di-iron centre per each (abc) monomer. Iron content determined by the Lovenberg colorimetric method [30], was 2 mol Fe per mol PHO dimer (abc) 2, lower than the theoretical value (4) of a di-iron centre per monomeric alpha subunit, but in accordance with experimental data obtained in similar conditions for MMO [42]. Sulfur was found to be absent. PHO secondary structure content was determined by circular dichroism (CD). The spectrum has the typical shape of mainly a-helical proteins, with a positive peak at 192 nm and two negative peaks at 209 and 222 nm. The prevalence of a-helices in the CD deduced secondary structure (99.6% as calculated with the CDNN deconvolution program [32]) confirmed the hypothesized structural homologies between PHO and MMO, whose crystallographic structure has been determined [19,20] and classified as all a-helical (SCOP: http://scop.mrc-lmb.cam.ac.uk/scop/) and mainly a-helical (CATH: http://www.biochem.ucl.ac.uk/bsm/cath_new/ index.html). Catalytic properties of purified PHO PHO oxygenase activity was detected by using a Clark electrode to evaluate oxygen consumption rates and further confirmed by NADH specific consumption and HPLC- monitoring of substrate degradation (Table 2). The decay curve of phenol concentration as evaluated by HPLC gave a good fit to a bi-exponential function (data not shown), suggesting the presence of a faster and a slower component within the reaction of phenol hydroxylation (the latter is likely to be related to the product-dependent inactivation of the protein complex and/or to irreversible inactivation by peroxides formed during reaction as previously observed [16]). The faster component was employed for calculating a turnover number of 20 ± 4 min )1 . This value is lower but still in line with the turnover numbers determined by the polarographic assay, i.e. 32 ± 8 min )1 under the same conditions. The phenol-dependent NADH consumption assay gave a calculated turnover number of 33 ± 10 min )1 (Table 2). The latter value was obtained after subtracting a contribution to the basal NADH consumption of 14 ± 2 min )1 , due to an uncoupled activity of PHR alone upon phenolic substrates addition. The presence of PHR and PHO (in a reaction mixture containing NADH as electron donor) was not sufficient per se to reconstitute a specific activity (with any of the three tested methods) upon substrate addition. The presence of a third component of the redox complex was therefore required to restore PH activity. The purification and structural characterization of this component, named PHI, is reported [26]. Table 2. Tested substrates for specific PHO activity at 24 °C, pH 7.4 in presence of 100 m M MOPS/NaOH buffer and 1.68 m M NADH. All substrates were used at final concentration, 1 m M . Substrate Activity as evaluated by: Oxygen consumption Substrate decay (HPLC) NADH consumption (lmol O 2 Æmin )1 Æ lmol )1 PHO) % (lmol consumedÆ min )1 Ælmol )1 PHO) % (lmol NADH consumedÆ min )1 Ælmol )1 PHO) a % Phenol 32 ± 8 100 20 ± 4 100 33 ± 10 100 o-cresol 12 ± 6 38 14 ± 4 70 20 ± 7 60 m-cresol 11 ± 4 35 14 ± 3 70 20 ± 7 60 p-cresol 8.6 ± 4 27 12 ± 5 60 20 ± 6 60 3-chlorophenol 5.8 ± 3 18 11 ± 4 55 5 ± 4 15 4-chlorophenol 5.8 ± 4 18 12 ± 4 60 9 ± 4 27 3,4-dimetylphenol 18 ± 7 56 9.6 ± 5 48 28 ± 6 85 a- naphthol (72 ± 18) b – 14 ± 5 70 (50 ± 27) b – b-naphthol (198 ± 28) b – 13.6 ± 5 68 (26 ± 35) b – a Basal NADH consumption, due to the reductase component (PHR) alone upon substrate addition, was subtracted to all measured values except naphthols (basal activity: 14±2 lmol NADH consumedÆmin )1 Ælmol )1 PHO). b Data are affected by the high basal oxygen and NADH consumption (76–172 lmol NADH consumedÆmin )1 Ælmol )1 PHO). Fig. 2. The UV/vis absorbance spectrum of purified PHO. Spectra of holo- (thick line) and apo- (thin line) PHO in 50 m M Hepes/NaOH buffer, pH 7.0. The apo-PHO (lacking any detectable iron and acti- vity) was prepared as decribed [45]. Protein concentration was 2.3 l M dimer (abc) 2 for both samples. The e 280 nm was calculated to be 643 800 M )1 Æcm )1 . Inset: magnified visible spectrum (300–500 nm). Ó FEBS 2003 A. radioresistens S13 phenol hydroxylase (Eur. J. Biochem. 270) 2249 Three parameters that might affect the kinetics of the overall PH reaction (optimum temperature, pH and ionic strength) were then studied. For this purpose, the optimal PHR/PHI/PHO fixed ratio of 2/1/1 was maintained [26]. The results are summarized in Fig. 3. Phenol hydroxylase activity was maximal between 0.075–0.1 M of ionic strength (Fig. 3A), whereas, the maximum oxygen consumption was detected at pH 7.5. (Fig. 3B). Temperature dependence of the phenol hydroxylase activity showed a biphasic pattern (Fig. 3C), with a first peak of activity measured at 24 °C, followed by a decrease and a second smaller peak of oxygen consumption at 32 °C. This may be due to the multi- component nature of the enzyme, as 32 °C corresponds to the maximum activity of the reductase component (PHR), which shuttles electrons for PHO catalysis [25]. As shown in Table 2, PHO from A. radioresistens S13 showed specific activity over a broad-substrate range, including methylated and mono-chlorinated phenols. Highly hydrophobic molecules like a-andb-naphthol were also recognized as seen by monitoring the substrate decay by HPLC. In these cases, the oxygen and NADH consumption values are higher than the substrate consumption and this is likely to be due to uncoupling reactions leading to species such as superoxide, hydrogen peroxide and hydroxyl radicals. The oxygen consumption rates might also be impaired by the contribution of a highly uncoupled NADH consumption activity due to the sole PHR/naphthol mixture, as evaluated by the spectrophotometric assay (Table 2). In contrast, phenolics with poly-chloro and nitro substi- tuents were not recognized and/or stabilized by the PHO active site. Among the substrates tested, no PHO activity was measured on p-hydroxybenzoic acid, m-hydroxy- benzoic acid, 2,4-dinitrophenol, 2,4-dichlorophenol, 3,4-dichlorophenol, 2,4,5-trichlorophenol, 2,2¢-dihydroxy- biphenyl and L -tyrosine. Discussion The oxygenase moiety (PHO) of the multicomponent phenol hydroxylase from A. radioresistens S13 seems to be endowed of a catalytic surface characterized by a relevant hydrophobicity, as suggested by evaluating its activity in different conditions of ionic strength. In fact, the plot of the activity vs. the ionic strength in the range 0–0.1 M shows a sigmoidal shape (Fig. 3A) where the enzyme activity reaches the maximum between 0.075 and 0.1 M . This suggests that the oxygenase reaction is facilitated by conditions that favour hydrophobic interactions between the components of PH, and/or between its active site and the substrate. As the whole PH is a multicomponent system, the ionic strength can initially promote interactions between PHR, Fig. 3. pH activity in the reconstituted complex in vitro: definition of the optimal conditions. pH activity was detected by Clark electrode in a reaction mixture containing 1.68 m M NADH, 0.6 l M PHO, 0.6 l M PHI, 1.2 l M PHR with addition of a final 1 m M phenol concentration. (A) Ionic strength dependence of PH activity. The experimental data were obtained either in Mops/NaOH buffer pH 7.4 or in Hepes/ NaOH buffer pH 7.4 at 24 °C. The continuous line from 0–0.1 M is the fit to a sigmoid. The dotted line represents the residual scattering of fitted data. (B) pH dependence. The activity was measured at a fixed ionic strength of 0.119 M inGood’sbuffersat24°C. (C) Temperature dependence. pH activity was calculated from the oxygen consumption rates after correction with Guy–Lussac factors at various temperature (20–40 °C). The activity was monitored in a Mops/NaOH buffer sys- tem at pH 7.4 (for further details see Materials and methods). 2250 S. Divari et al. (Eur. J. Biochem. 270) Ó FEBS 2003 PHI and PHO and between PHO and substrate. A further increase in ionic strength at levels higher than 0.12 M could shield critical electrostatic interactions within the multi- enzymatic complex, thus acting as a disaggregating factor. As catechol 1,2-dioxygenase, the enzyme that acts subse- quently to PH in the detoxification cascade, has already been demonstrated to possess a particular surface hydro- phobicity [43], a further elucidation of the hydrophobic surface interactions of both enzymes will be pursued. In vivo PHO and catechol 1,2-dioxygenase might form a stable complex or attain transient but very efficient interactions that would override the problems of two soluble proteins that catalyze sequential reactions without being anchored to a membrane system (as is instead the case for the best- known redox chains, i.e. respiration and photosynthesis). Analysis of the amino acid sequence of the cloned catalytic PHO a-subunit further emphasize the crucial role of hydrophobic interactions occurring in the active site cleft for substrate recognition. Alignment of the sequence of the a-subunit with that of methane monoxygenase (Table 1), the closest PH related system whose 3D structure has been elucidated [19,20], allowed the identification of the consen- sus sequences for the coordination of a dinuclear oxo-iron centre. The presence of this kind of nonhaem iron is also suggested by PHO UV/vis spectral features in the visible region (Fig. 2). Similar spectra have been observed recently for the phenol hydroxylase from Pseudomonas sp. strain CF600 [16]. Conserved hydrophobic amino-acids are also present in the a-subunit of PHO, corresponding to the second coordination sphere of the prosthetic metal. These are known to define the substrate recognition pocket in well characterized monooxygenases similar to PHO (i.e. MMO and AMO [35,36]). Moreover, a high percentage of identity was identified between the PHO a-subunit of A. radioresis- tens S13 and that of PHO4 of A. calcoaceticus NCBI 8250 and P3 from Pseudomonas sp. strain CF600 [15]. For the latter two enzymes, no correlations are reported between the substrate specificity and the role of conserved hydrophobic residues. The functional characterization of PHO and the analysis of its activity on other potential aromatic substrates allow us to propose a model to explain the fine modulation of substrate recognition. PHO is apparently able to recognize bulkier substrates with higher hydrophobicity than phenol rings, such as cresols, monochlorophenols and naphthols (Table 2). A marked hydrophobicity of the active site could also be responsible for the null activity on phenolics and aromatics with strongly hydrophilic or charged substituents, such as benzoic acid and tyrosine, that would not be easily stabilized in the vicinity of the catalytic site. The experi- mental data on relaxed substrate specificity of PHO suggest that the active site arrangement of conserved hydrophobic amino-acids might result, in this case, in a larger or more flexible active site pocket. The structural basis of the fine tuning of PHO-substrate recognition might be due to substitutions of critical hydrophobic residues, such as Ala117, Phe236 and Ile239 occurring in the MMO from M. capsulatus, with smaller homologous amino acids (namely glycine for the first two cases and alanine for the last) that would cause an easier accessibility to the dinuclear oxo-iron centre (Table 1). These particular substitutions are also observed in the two available sequences of multicom- ponent phenol hydroxylases [11,12]. On the other hand, both in PHs and in similar enzymes that hydroxylate aromatics [6,37,38] a phenylalanine is present in a position corresponding to a glycine (Gly208) in MMO (Table 1). The latter is the conserved hydrophobic residue adjacent to a glutamic acid coordinated to the iron. The closeness to this first sphere coordinating residue suggests that the Phe aromatic ring might be involved in ÔsandwichingÕ the phenolic substrate in a correct orientation for catalysis. Although one flavin dependent phenol hydroxylase was reported to work efficiently with methyl- and cloro-phenols [44], the di-iron centre can potentially support the hydroxy- lation of nonactivated molecules such as benzene, toluene, xylene. The versatility in substrate recognition of the PHO from A. radioresistens S13 is of particular interest and makes this enzyme a good candidate for future biotechno- logical exploitation, even if the in vitro PH assay demon- strated that the enzyme activity is affected by a certain degree of uncoupling. This is due possibly to the production of peroxide species during the oxygenating process. Also, an increased NADH consumption by the reductase component might indicate a partial denaturation of PHR and/or alteration of its redox centre. This is particularly important for naphthols. HPLC data indicate nonetheless that the various substrates are recognized and consumed by the enzyme. The accumulation of hydroxylated products such as catechol, chlorocatechol and methylated catechols, as well as naphthols (as identified by their retention times and spectra) was observed during HPLC assay of substrate consumption, although their low levels affect the data scattering (data not shown). Further characterization of products will be pursued in order to evaluate the degree of recognition and processing of substrates other than phenol. The biphasic nature of phenol disappearance, as moni- tored by HPLC, suggests that accumulation of the end product might inactivate the enzyme, and therefore the presence of the downstream enzyme, catechol 1,2 dioxy- genase (C1,2O) is necessary to shift the equilibrium towards the detoxification reactions. 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(2000) Purification and catalytic properties of two catechol 1,2- dioxygenase isozymes from benzoate-grown cells of Acinetobacter radioresistens. J. Protein Chem. 19, 709–716. 44. Neujahr, K.J. & Kjellen, K.G. (1978) Phenol hydroxylase from yeast. Reaction with phenol derivatives. J. Biol. Chem. 253, 8835–8841. 45. Smith, D.D.S. & Dalton, H. (1992) Evidence for two histidine ligands at the diiron site of methane monooxygenase. Eur. J. Biochem. 210, 629–633. Ó FEBS 2003 A. radioresistens S13 phenol hydroxylase (Eur. J. Biochem. 270) 2253 . wastes, from which the strain was isolated by our group [27]. This paper describes the purification, the characterization and the catalytic properties of the oxygenase component of PH from A. radioresistens. 2,4,5-trichlorophenol, 2,2¢-dihydroxy- biphenyl and L -tyrosine. Discussion The oxygenase moiety (PHO) of the multicomponent phenol hydroxylase from A. radioresistens S13 seems to be endowed of a catalytic. The oxygenase component of phenol hydroxylase from Acinetobacter radioresistens S13 Sara Divari 1 , Francesca Valetti 1 , Patrizia Caposio 4 ,