the oxygenase component of phenol hydroxylase from acinetobacter radioresistens s13

The oxygenase component of phenol hydroxylase fromAcinetobacter radioresistens S13

Sara Divari1, Francesca Valetti1, Patrizia Caposio4, Enrica Pessione1, Maria Cavaletto2, Ersilia Griva1,Giorgio Gribaudo4, Gianfranco Gilardi1,3 and Carlo Giunta1

1Dipartimento di Biologia Animale e dell’Uomo, Universita di Torino, Italy; 2Dipartimento di Scienze e Tecnologie Avanzate,

Universita del Piemonte Orientale, Alessandria, Italy; 3Department of Biological Sciences, Imperial College of Science,

Technology and Medicine, London, UK; 4Dipartimento di Sanita Pubblica e Microbiologia, Universita di Torino, Italy

Phenol hydroxylase (PH) from Acinetobacter radioresis-tens S13 represents an example ofmulticomponent aromaticring 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 molecularoxygen and catalyse the mono-hydroxylation of substrates(phenol, and with less efficiency, chloro- and methyl-phenoland naphthol). PHOwas purified from extracts ofA. radio-resistens S13 cells and shown tobe adimerof 206 kDa.Eachmonomer is composed by three subunits: a (54 kDa), b(38 kDa) andc (11 kDa).The gene encodingPHOa (namedmopN) was cloned and sequenced and the correspondingamino acid sequence matched with that of functionallyrelated oxygenases. By structural alignment with the cata-lytic subunits of methane monooxygenase (MMO) andalkene monooxygenase, we propose that PHO a containsthe enzyme active site, harbouring a dinuclear iron centre

Fe-O-Fe, as also suggested by spectral analysis. Conservedhydrophobic amino acids known to define the substraterecognition pocket, are also present in the a-subunit. Theprevalence of a-helices (99.6%) as studied by CD confirmedthe hypothized structural homologies between PHO andMMO. Three parameters (optimum ionic strength, tem-perature and pH) that affect kinetics of the overall phenolhydroxylase reaction were further analyzed with a fixedoptimal PHR/PHI/PHO ratio of 2/1/1. The highest levelof activity was evaluated between 0.075 and 0.1 M of ionicstrength, the temperature dependence showed a maximumof activity at 24 �C and finally the pH for optimal activitywas determined to be 7.5.

Keywords: multicomponent monooxygenase; phenolhydroxylase; purification; molecular cloning; catalyticsubunit.

Phenol-degrading aerobic bacteria are able to convertphenol into nontoxic intermediates of the tricarboxylic acidcycle via an ortho or meta pathway [1]. The first step ofboth routes is the monohydroxylation of the ortho positionof the aromatic ring [2]. The enzyme responsible for thisreaction is the monooxygenase phenol hydroxylase (PH).Aromatic monooxygenases are divided into two groups:

activated-ring monooxygenases (monocomponent) andnonactivated-ring enzymes (multicomponent). In the lattercase, the active site must contain a strong hydroxyl-generating-unit, i.e. a dinuclear iron centre in which anoxygen 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 tosupply electrons from NAD(P)H to the dinuclear ironcentre itself. Such a multicomponent organization is presentin a number of enzymes that are able to hydroxylate andstart the detoxification process of poorly reactive aromaticsand aliphatics, often recalcitrant to degradation. Amongthese molecules examples are toluene, that is converted top-hydroxytoluene by toluene-4-mono-oxygenase in Pseudo-monas mendocinaKR1 [5]; xylene, the substrate of a xylene/toluene monooxygenase in Pseudomonas stutzeri OX1 [6];methane, that is converted to methanol by methanemonooxygenases 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: [email protected]

Abbreviations: AMO, alkene monooxygenase; AMOa, alkene mono-

oxygenase a subunit fromNocardia corallinaB-276;AMO Py2, alkene

monooxygenase from Xanthobacter Py2; C1,2O, catechol 1,2 di-

oxygenase; MMOM, methane monooxygenase; MMO B, methane

monooxygenase fromMethylococcus capsulatus Bath; MMOMz,

M methane monooxygenase fromMethylosinus trichosporium OB3b;

PH, phenol hydroxylase; PHI, phenol hydroxylase regulatory protein;

PHR, phenol hydroxylase reductase; PHO, phenol hydroxylase

oxygenase; T3MO, toluene-3-monooxygenase from Pseudomonas

pickettiiPKO1; T4MO, toluene-4-monooxygenase fromPseudomonas

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) � FEBS 2003 doi:10.1046/j.1432-1033.2003.03592.x

Methylosinus thrichosporium [8] and Methylocystis [9];alkenes, converted to the corresponding epoxides by thealkenes monooxigenase of Nocardia corallina B-276 [10].Surprisingly, in a few bacterial strains [11,12], a very similarsystemwas also described to recognize phenol, even if in thiscase the aromatic ring to be processed is activated by thehydroxyl group and a simple FAD-dependent monocom-ponent enzyme would be able to catalyze the reaction, asreported for Pseudomonas pickettii [13] and Bacillus stearo-thermophylus [14]. The multimeric phenol hydroxylase fromPseudomonas sp. strain CF 600 [1,15,16] is of particularinterest because its biochemical and genetic characteri-zation indicated an organization very similar to bothmethane monooxygenase from methanotrophs [17–20]and to alkene monooxygenase from Nocardia corallina[21]. All these monooxygenases are composed of oneNADH binding monomeric reductase, one multimeric(abc)2 oxygenase containing the dinuclear iron-centre (onthe a subunit) and binding oxygen and substrate, plus asmall regulatory component whose NMR structure hasbeen reported recently [22–24]. Among the Acinetobactergenus, only genetic data are available for the phenol-degrading A. calcoaceticus NCIB 8250 [12]. These show agenomic organization of the PH encoding genes very similarto that reported for P. sp. strain CF600 in addition to asignificant amino acid sequence homology. Therefore thereare at least two known PHs expressed by different bacterialgenera with a multicomponent organization.In a previous paper, we showed that PH from Acineto-

bacter radioresistens LMG S-13648 (thereafter, designed toas A. radioresistens S13) consists of three components [25].The reductase component, PHR, is a monomeric iron-sulfur-flavoprotein [25] that transfers reducing equivalentsfrom NAD(P)H to the oxygenating moiety PHO describedin this work, and PHI is an intermediate componentnecessary for catalysis [26]. This multicomponent enzymeconfers to the organism a very high (100 mgÆL)1Æ h)1)phenol degradation rate when this substrate is the solecarbon and energy source. The potential of this organism inbioremediation is shown by its ability to grow on activatedsludges of industrial wastes, from which the strain wasisolated by our group [27].This paper describes the purification, the characterization

and the catalytic properties of the oxygenase component ofPH from A. radioresistens S13, as well as the molecularcloning 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 andHowell [28] minimal medium in which phenol was the solecarbon source. Phenol was added in a fed-batch fermenta-tion procedure (100 mgÆL)1Æh)1) and the culture wasincubated at 30 �C for 23–24 h. Cells were harvested byultracentrifugation at 15 000 g, washed twice in 50 mLHepes/NaOH buffer, pH 7.0 and stored frozen ()80 �C).Biomass (200 g per 200 mL) in 50 mM Hepes/NaOHbuffer, pH 7.0 were sonicated on ice for a total time of20 min at 20 kHz with intervals of 1 min (MicrosonixSonicator Ultrasonic Liquid Processor XL2020). The

obtained soluble extracts were then centrifuged at 100 000g for 1 h at 4 �C (ultracentrifuge LB60M Beckman). Thesupernatants were considered as the crude extracts.

Phenol hydroxylase assay

PH activity was measured polarographically by means of aClark-type electrode (YSI Model 5300). The assay wascarried out in the presence of 1.68 mM NADH, 0.6 lMPHO, 1.2 lM PHR, 0.6 lM PHI and 100 mMMops/NaOHbuffer, pH 7.4 at 24 �C. The reaction was started by adding1 mM phenol (Fluka). The same experiment was performedusing the following substrates (1 mM): 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. Poorlywater soluble compounds were prepared as 300-fold con-centrated stock solutions in methanol and only 10 lL wereadded to the reaction mixture. No interference or proteindamage was observed due to the presence of this amount ofmethanol. Kinetics of phenol consumption by PH reconsti-tuted complex were also evaluated by a discontinuous assayat 24 �C, in 100 mM Mops/NaOH buffer pH 7.4 at thesame reactant concentration used for the oxygen consump-tion assay (1.68 mM NADH, 0.6 lM PHO, 1.2 lM PHR,0.6 lM PHI and 1 mM phenol). Residual phenol concen-tration at different times of reaction was evaluated byHPLC (Merck Hitachi L6200 system) on a LichrosphereRP-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 wasperformed in order to separate the phenol peak that wasmonitored at 270 nm by a Diode Array detector system(Merck Hitachi L4500) and quantitated by the correctingpeak area against the total area. The obtained data werefitted to a biexponential decay function and only the fastercomponent was considered for turnover number calcula-tion. The same procedure was applied to the other testedsubstrates, adjusting HPLC eluents to pH ¼ 3.0 withconcentrated H2SO4, and varying the monitored wave-length to the corresponding k-max. A third kind of assaywas based on the continuous measurement of NADHconsumption. The same reaction mixture described abovewas monitored at 24 �C for NADH disappearance at340 nmwith a BeckmanDU 70 spectrophotometer and thekinetics calculated by taking the initial velocity as thetangent to the obtained curve. The reaction was started bysubstrate 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) equilibratedpreviously with 50 mM Hepes/NaOH buffer pH 7.0. Theelution was obtained with a 0–0.5 M sodium sulphategradient in 50 mM Hepes/NaOH buffer pH 7.0 (finalvolume, 1.1 L). After ultrafiltration (membrane Diaflo,cut off 30 kDa; Amicon), the PHO containing fractionswere applied on a Q-Sepharose FF (Pharmacia) column(1.3 · 26 cm) equilibrated with 50 mM Hepes/NaOH

� FEBS 2003 A. radioresistens S13 phenol hydroxylase (Eur. J. Biochem. 270) 2245

buffer, pH 7.0 containing 0.15 M sodium chloride. PHOwas eluted from this column with a 0.15–0.35 M NaCl lineargradient (total volume 1.2 L) in 50 mM Hepes/NaOHbuffer pH 7.0. Active fractions eluted were concentratedby ultrafiltration and applied on a hydrophobic Phenyl-Sepharose 6FF column (Pharmacia) (1.3 · 13 cm). Thecolumn was equilibrated with 50 mM Hepes/NaOH bufferpH 7.0, containing 0.15 M sodium sulphate and the sampleadjusted to the corresponding ionic strength with the samebuffer. A mixed gradient was applied with an initial linearsection from 0.15–0 M Na2SO4 in 50 mM Hepes/NaOHbuffer pH 7.0 (50 mL), an isocratic step of 120 mL of50 mM Hepes/NaOH buffer pH 7.0, a 50-mL linear gradi-ent from 50 mMHepes/NaOHbuffer pH 7.0 to water and afinal isocratic step in 100% water. PHO was eluted in thislast section of the mixed gradient. In order to avoidprolonged exposure of PHO to water, the 3 mL elutedfractions were collected in tubes containing the same volumeof 200 mM Hepes/NaOH buffer pH 7.0. PHO fractions in100 mMHepes/NaOH buffer, pH 7.0 were concentrated, asdescribed previously, and stored at )80 �C until required.An average yield of 0.1 lmol of purified protein wasobtained from 200 g of biomass following such procedure.

Characterization

Total protein concentration was estimated colorimetricallyby the Bradford method [29] using BSA as standard.Molecular mass was determined by gel filtration andSDS/PAGE. A Superdex 200-FPLC (Pharmacia) column(1.6 · 60 cm) was equilibrated with 50 mM Hepes/NaOHbuffer pH 7.0 with 0.05 M Na2SO4 and calibrated withthyroglobulin (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 determinedby 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 protomerswas achieved by RP-HPLC. PHO (32 lM) was mixed to178 lL of 50% acetonitrile solution and 1% formic acid.Protomers generated were purified using a HPLC Merck-Hitachi L6200 system equipped with a Lichosphere 100RP-8 column (Merck). The flow rate was 1 mLÆmin)1. Thecolumn was equilibrated with solvent A (water/trifluoro-acetic acid, 100/0.08) and protomers were eluted using alinear gradient of 20–100% solvent B (water/acetonitrile/trifluoroacetic acid, 10/90/0.08) over 80 min. N-Terminalsequences 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 sequencedfollowing the Edman degradation, using a 470-A phasesequencer (Applied Biosystems USA). The isoelectric pointwas determined by analytical IEF electrophoresis (Phast-System, Pharmacia); the markers were those supplied byPharmacia (pI calibration kit). The iron and sulfur contentwas determined by colorimetric methods as describedpreviously [30,31]. CD measurements were performed ona 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 pathlength cell and a scan speed of 20 nmÆmin)1 with a responsetime of 8 s. The CD spectrum reported is the average ofthree scans at 0.5 nm resolution and a bandwidth of 2 nm.The concentration was 1.25 lM for PHO. The spectrumwasanalyzed with the CDNN deconvolution program [32].

Construction and screening of the genomic library

The A. radioresistens S13 genomic library was prepared asdescribed [33]. Briefly, purified genomic DNA (10 lg) waspartially digested with Sau3AI, and DNA fragments(10–20 kb) were fractionated by preparative gel electro-phoresis. Thereafter, the partially digested genomic DNAwas partly completed with the Klenow fragment of E. coliDNA polymerase I and dGTP and dATP, and then ligatedto LambdaGEM-12 XhoI half-site arms (Promega) accord-ing to the manufacturer’s instructions. TheA. radioresistensS13 genomic library resulted in 1.3 · 105 independentrecombinant clones that were stored and screened with-out amplification. It was plated on layers of susceptiblebacteria host at a density of 5000 plaques per 150-mmdiameter Petri dish, and a duplicate set of nylon filters(Hybond-N+, Amersham) was taken from each filter.Filters were hybridized with a radiolabeled DNA probein a solution containing 5 · NaCl/Cit-5 · Denhardt’s-1%SDS-100 lgÆmL)1 of denatured salmon sperm DNA for18 h at 65 �C. Filters were then washed twice for 20 min at65 �C in 2 · NaCl/0.1%Cit SDS and then twice for 20 minat 65 �C in 0.2 · NaCl/Cit-0.1% SDS. DNA inserts fromrecombinant phages of interest were prepared, restrictedwith EcoRI and subcloned into pGEM-7Zf(+) (Promega).DNA templates were then sequenced by the dideoxy-chainterminator method using a Sequenase 2.0 DNA sequencingkit (USB). Sequence data were used for homology search inthe GenBank database and have been deposited in the samedatabase with accession number for AF521658.The hybridization probe was prepared by means of

PCR using degenerated primers synthesized accordingto the amino acid sequences, SQVKTTVKKL andLLSIVAMMM. PCR was carried out with 100 ng ofA. radioresistens S13 genomic DNA in PCR buffer (10 mMTris/HCl, pH 8.3, 50 mM KCl, 1.5 mM MgCl2) containingthe two primers (0.250 lM each), the four dNTPs (200 lMeach) and 2.5 U of Taq DNA polymerase (Perkin-Elmer).Samples were subjected to 30 cycles of amplification withthe following cycle profile: denaturation at 94 �C for 1 min,annealing to 50 �C for 1 min, extension at 72 �C for 2 min.

pH, temperature and ionic strength dependenceof enzyme activity

PHO activity was determined at 24 �C in the pH range5.5–9, using Good’s buffers (50 mM Mes-Mops-CHES)adjusted at a fixed ionic strength of 0.119 M by addition,where necessary, ofNaCl. No difference was observed in thereaction upon adjusting ionic strength by buffer concentra-tion or by addition of NaCl. The activity was evaluated asoxygen consumption by a Clark electrode. The temperaturedependence of phenol hydroxylation reaction at pH 7.4 wasinvestigated 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) � FEBS 2003

employed for calculating the saturating oxygen concentra-tion, at the tested temperature, in the aqueous reactionmixture. Ionic strength dependence of PH activity wasstudied in the same reaction condition described above, at24 �C and pH 7.4. The ionic strength was varied from4–164 mM by adjusting the concentration of Mops/NaOHbuffer in a range 10–250 mM. The experiment was repeatedin Hepes/NaOH buffer pH 7.4 in the same concentrationrange. The precise ionic strength at each buffer concentra-tion was calculated by means of the on-line programavailableathttp://www.bi.umist.ac.uk/users/mjfrbn/buffers/makebuf.asp [34].Dataobtained fromthe twobuffer systemswere found coincident within the experimental error andtherefore gathered in the same dataset. Perfectly comparableresults were also obtained when the ionic strength wasadjusted to the desired value byNaCl addition.

Results

PHO purification

Purification of PHO was achieved by two anion exchangecolumns followed by a hydrophobic interaction chroma-tography.

The molecular mass value of the purified PHO wasestimated to be 207 kDa by gel-permeation, whilst threebands of 54 kDa, 37.8 kDa and 11.6 kDawere observed onSDS/PAGE gels (Fig. 1A). This suggests a hexamericstructure such as (abc)2. The three subunits were alsoseparated by RP-HPLC with a weaker interaction ofc subunit to the C8 column matrix and a higher affinityof the b and a subunits (Fig. 1B). The isoelectric point of thepurified PHO was determined to be 6.74.

N-terminal primary structure alignment

N-terminal sequences of the three subunits of PHO weredetermined to be SQVKTTVKKL for the a-, TLEIKTAGIE for the b- and SVRAIRPDYD for the c-subunit.These sequences were compared with analogous sequencesof PHOs extracted from different bacterial species thatexpress multicomponent phenol hydroxylases. The degreeof identity is very high (60%) for the a- and b-subunitsof A. radioresistens S13 and A. calcoaceticus [12], lower(40–20%) between A. radioresistens S13 and Pseudomonassp. strain CF600 [11] and very low (30–20%) amongthe 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 are 54 kDa, 38 kDa and 11 kDa,respectively. (B) PHO protomer separation by

RP-HPLC. PHO (32 lM) was mixed to178 lL of 50%H2O-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

1 mLÆmin)1. The column was equilibratedwith solvent A (H2O/Trifluoroacetic acid,

100/0.08) and protomers were eluted using a

linear gradient of 20–100% solvent B (H2O/

CH3CN/trifluoroacetic acid, 10/90/0.08) over

80 min.


Cloning and nucleotide sequence of the phenolhydroxylase PHO a-subunit

The gene encoding for the PHO a-subunit was obtainedfrom a genomic library of A. radioresistens S13, preparedand screened with a specific DNA probe obtained by PCRamplification of genomic DNA with degenerated oligo-nucleotides designed on the NH2-terminal sequence and onpeptides derived by trypsin digestion of the purified protein.After subcloning and sequence analysis, the nucleotide anddeduced amino acid sequences were matched with proteinsequences. To complete the cloning of the gene of the PHOa-subunit, about 104 phage plaques were screened with thePCR product as probe and the P19.2 clone was selected forPHO a-encoding sequences. The gene encoding for thePHO a-subunit (mopN) was located on a 1.9-kb EcoRIsegment of the P19.2 clone and the nucleotide sequence wasdetermined. The sequence of the 1.9 kb EcoRI fragmentrevealed that the mopN gene of A. radioresistens S13encodes an ORF of 1527 base pairs corresponding to apolypeptide of 509 amino acids with a calculated molecularweight of 59.9 kDa. This value is in agreement (within theexperimental error) with that obtained by SDS/PAGEresolution of purified PHO subunits. Further analysis(Table 1) of the whole deduced amino acid sequencesrevealed 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 coordinationof the catalytic dinuclear iron centre, as already described inMMO from M. capsulatus and M. trichosporium and inPHOs from Pseudomonas CF 600 and A. calcoaceticusNCIB8250 (Table 1). Furthermore, many hydrophobicresidues of the protein aligned with conserved amino acidsof MMO active site pocket (Table 1). PHO a shows thehighest degree of identity (92%) with the DMS oxygenasecomponent of A. sp. strain 20B (accession numberBAA2333.1) [39] and with the PHO component 4 (ORF 4)of A. calcoaceticus NCBI 8250 (accession numberCAA85383.1) [12]. High similarity (70% identity) was alsoobserved with both the PHO component D of P. putidastrain H (accession number CAA56743.1) [40], and the P3protein 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 tolueneortho-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 peakat 280 nm (Fig. 2). The e280 nm was calculated to be643 800 M)1Æcm)1 in 50 mM Hepes/NaOH buffer, pH 7.0.In the visible region, a broad shoulder is highlighted whencomparing holo- and apo-PHO (Fig. 2, inset). This can beattributed to the presence of oxo-bridged di-iron centres asreported recently [16] for phenol hydroxylase from P. sp.strain CF600. For the latter, the extinction coefficient at350 is between 4800 and 6000 M)1Æcm)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, close to T

able

1.Sequence

alignmentofA.radioresistens

S13PHO

awiththeactivesite

andmetalcoordinatingregionsofhomologousdi-nuclearoxo-ironoxygenases.Conservedresiduesinvolvedintheoxo-iron

centrecoordinationareindicatedinbold.Hydrophobicresiduesdelimitingtheactivesitecavityareunderlined.Residuesarenumberedonlyfortheenzymeswithknown3D

structures[19,20].PHO

a,A.radioresistensS13phenolhydroxylase

acomponent;MMOB,M.capsulatusmethanemonooxygenase[17];MMOM,M.trichosporiummethanemonooxygenase[18];AMO

a,N.corallinaalkene

monoxygenase;AMOPy2,X.Py2alkenemonoxygenase[36];Xyl/TMO,P.stutzerixylene/toluenemonooxygenase[6];T4MO,P.mendocinatoluene4monooxygenase[37];T3MO,P.pickettiitoluene3

monooxygenase[38].

Enzyme

Residue

Sequence

Residue

Sequence

Residue

Sequence

Residue

Sequence

Residue

PHO

a–

LEYQAFQG

–/–

MQSIDELRHV

–/–

FEFLLAISFAFEYVLTNLLFV

–/–

TFGFSAQSDEARHMTLG

||

:::

|:||:||

:|

|::::|::|||:|

|:::|:::||:|||

|

MmoB

110

LEVGEYNA

117/139

AQVLDEIRHT

148/198

VECSLNLQLVGEAGFTNPLIV

218/234

TVFLSIETDELRHMANG

250

MmoM

110

GEYNAIAA

117/139

AQVLDEIRHT

148/198

VECSVNLQLVGDTCFTNPLIV

218/234

TVFLSVRTDELRHMANG

250

Amo

a–

LTNAEYQA

–/–

AQMLDEVRHA

–/–

LDVIIDLNIVAETAFTNILLV

–/–

SVFLSIQSDEARHMANG

AmoPy2

–VEHMAVTM

–/–

FGMLDETRHT

–/–

VEAALATSLTLEHGFTNIQFV

–/–

NLLSSIQTDEARHAQLG

Xyl/TMO

–EEYAASTA

–/–

FGMMDENRHG

–/–

VAVSIMLTFAFETGFTNMQFL

–/–

SLISSIQTDESRHAQQG

T4MO

–GEYAAVTG

–/–

FGMMDELRHG

–/–

ISVAIMLTFSFETGFTNMQFL

–/–

NLISSIQTDESRHAQQG

T3MO

–GEYAAMSA

–/–

FGMLDENRHG

–/–

IDIAIMLTFAFETGFTNMQFL

–/–

SLISSIQTDESRHAQIG


the expected value of one di-iron centre per each (abc)monomer. Iron content determined by the Lovenbergcolorimetric method [30], was 2 mol Fe per mol PHOdimer (abc)2, lower than the theoretical value (4) of a di-ironcentre permonomeric alpha subunit, but in accordance withexperimental 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 shapeof mainly a-helical proteins, with a positive peak at 192 nmand two negative peaks at 209 and 222 nm. The prevalenceof a-helices in the CD deduced secondary structure (99.6%

as calculated with the CDNN deconvolution program [32])confirmed the hypothesized structural homologies betweenPHO andMMO, whose crystallographic structure has beendetermined [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 Clarkelectrode to evaluate oxygen consumption rates and furtherconfirmed by NADH specific consumption and HPLC-monitoring of substrate degradation (Table 2). The decaycurve of phenol concentration as evaluated by HPLC gavea good fit to a bi-exponential function (data not shown),suggesting the presence of a faster and a slower componentwithin the reaction of phenol hydroxylation (the latter islikely to be related to the product-dependent inactivation ofthe protein complex and/or to irreversible inactivation byperoxides formed during reaction as previously observed[16]). The faster component was employed for calculating aturnover number of 20 ± 4 min)1. This value is lower butstill in line with the turnover numbers determined by thepolarographic assay, i.e. 32 ± 8 min)1 under the sameconditions. The phenol-dependent NADH consumptionassay gave a calculated turnover number of 33 ± 10 min)1

(Table 2). The latter value was obtained after subtractinga contribution to the basal NADH consumption of14 ± 2 min)1, due to an uncoupled activity of PHR aloneupon phenolic substrates addition.The presence of PHR and PHO (in a reaction mixture

containing NADH as electron donor) was not sufficient perse to reconstitute a specific activity (with any of the threetested methods) upon substrate addition. The presence of athird component of the redox complex was thereforerequired to restore PH activity. The purification andstructural 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 mM MOPS/NaOH buffer and 1.68 mM NADH. All

substrates were used at final concentration, 1 mM.

Substrate

Activity as evaluated by:

Oxygen consumption Substrate decay (HPLC) NADH consumption

(lmol O2Æmin)1Æ

lmol)1 PHO) %

(lmol consumedÆmin)1Ælmol)1 PHO) %

(lmol NADH consumedÆmin)1Ælmol)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)1Ælmol)1PHO). b Data are affected by the high basal oxygen andNADH consumption (76–172 lmol NADH consumedÆmin)1Ælmol)1 PHO).

Fig. 2. The UV/vis absorbance spectrum of purified PHO. Spectra of

holo- (thick line) and apo- (thin line) PHO in 50 mM 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 lMdimer (abc)2 for both samples. The e280 nm was calculated to be643 800 M)1Æcm)1. Inset: magnified visible spectrum (300–500 nm).


Three parameters that might affect the kinetics of theoverall PH reaction (optimum temperature, pH and ionicstrength) were then studied. For this purpose, the optimalPHR/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 wasdetected at pH 7.5. (Fig. 3B). Temperature dependence ofthe 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 oxygenconsumption at 32 �C. This may be due to the multi-component nature of the enzyme, as 32 �C corresponds tothe 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,includingmethylatedandmono-chlorinatedphenols.Highlyhydrophobic molecules like a- and b-naphthol were alsorecognized as seen by monitoring the substrate decay byHPLC. In these cases, the oxygen andNADH consumptionvalues are higher than the substrate consumption and this islikely to be due to uncoupling reactions leading to speciessuch as superoxide, hydrogen peroxide and hydroxylradicals. The oxygen consumption rates might also beimpaired by the contribution of a highly uncoupled NADHconsumptionactivity due to the solePHR/naphtholmixture,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 PHOactive site. Among the substrates tested, no PHO activitywas 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 multicomponentphenol hydroxylase from A. radioresistens S13 seems to beendowed of a catalytic surface characterized by a relevanthydrophobicity, as suggested by evaluating its activity indifferent conditions of ionic strength. In fact, the plot of theactivity vs. the ionic strength in the range 0–0.1 M shows asigmoidal shape (Fig. 3A)where the enzyme activity reachesthe maximum between 0.075 and 0.1 M. This suggests thatthe oxygenase reaction is facilitated by conditions thatfavour hydrophobic interactions between the componentsof PH, and/or between its active site and the substrate. Asthe whole PH is a multicomponent system, the ionicstrength 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 mM NADH, 0.6 lM PHO, 0.6 lMPHI, 1.2 lM PHRwith addition of a final 1 mM 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/

NaOHbuffer pH 7.4 at 24 �C. The continuous line from 0–0.1 M is thefit 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 in Good’s buffers at 24 �C. (C) Temperaturedependence. 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).


PHI and PHO and between PHO and substrate. A furtherincrease in ionic strength at levels higher than 0.12 M couldshield 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 alreadybeen demonstrated to possess a particular surface hydro-phobicity [43], a further elucidation of the hydrophobicsurface interactions of both enzymes will be pursued. In vivoPHO and catechol 1,2-dioxygenase might form a stablecomplex or attain transient but very efficient interactionsthat would override the problems of two soluble proteinsthat catalyze sequential reactions without being anchored toa 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 roleof hydrophobic interactions occurring in the active site cleftfor substrate recognition. Alignment of the sequence of thea-subunit with that of methane monoxygenase (Table 1),the closest PH related system whose 3D structure has beenelucidated [19,20], allowed the identification of the consen-sus sequences for the coordination of a dinuclear oxo-ironcentre. The presence of this kind of nonhaem iron is alsosuggested by PHO UV/vis spectral features in the visibleregion (Fig. 2). Similar spectra have been observed recentlyfor the phenol hydroxylase from Pseudomonas sp. strainCF600 [16]. Conserved hydrophobic amino-acids are alsopresent in the a-subunit of PHO, corresponding to thesecond coordination sphere of the prosthetic metal. Theseare known to define the substrate recognition pocket in wellcharacterized monooxygenases similar to PHO (i.e. MMOand AMO [35,36]). Moreover, a high percentage of identitywas identified between the PHO a-subunit of A. radioresis-tens S13 and that of PHO4 of A. calcoaceticus NCBI 8250and P3 from Pseudomonas sp. strain CF600 [15]. For thelatter two enzymes, no correlations are reported between thesubstrate specificity and the role of conserved hydrophobicresidues.The functional characterization of PHO and the analysis

of its activity on other potential aromatic substrates allowus to propose a model to explain the fine modulation ofsubstrate recognition. PHO is apparently able to recognizebulkier substrates with higher hydrophobicity than phenolrings, such as cresols, monochlorophenols and naphthols(Table 2). A marked hydrophobicity of the active site couldalso be responsible for the null activity on phenolics andaromatics with strongly hydrophilic or charged substituents,such as benzoic acid and tyrosine, that would not be easilystabilized in the vicinity of the catalytic site. The experi-mental data on relaxed substrate specificity of PHO suggestthat the active site arrangement of conserved hydrophobicamino-acids might result, in this case, in a larger or moreflexible active site pocket. The structural basis of the finetuning of PHO-substrate recognition might be due tosubstitutions of critical hydrophobic residues, such asAla117, Phe236 and Ile239 occurring in the MMO fromM. capsulatus, with smaller homologous amino acids(namely glycine for the first two cases and alanine for thelast) that would cause an easier accessibility to the dinuclearoxo-iron centre (Table 1). These particular substitutions arealso 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 hydroxylatearomatics [6,37,38] a phenylalanine is present in a positioncorresponding to a glycine (Gly208) in MMO (Table 1).The latter is the conserved hydrophobic residue adjacent toa glutamic acid coordinated to the iron. The closeness to thisfirst sphere coordinating residue suggests that the Phearomatic ring might be involved in �sandwiching� thephenolic 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 PHOfrom A. radioresistens S13 is of particular interest andmakes 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 certaindegree of uncoupling. This is due possibly to the productionof peroxide species during the oxygenating process. Also, anincreasedNADHconsumption by the reductase componentmight indicate a partial denaturation of PHR and/oralteration of its redox centre. This is particularly importantfor naphthols. HPLC data indicate nonetheless that thevarious substrates are recognized and consumed by theenzyme. The accumulation of hydroxylated products suchas catechol, chlorocatechol and methylated catechols, aswell as naphthols (as identified by their retention times andspectra) was observed during HPLC assay of substrateconsumption, although their low levels affect the datascattering (data not shown). Further characterization ofproducts will be pursued in order to evaluate the degree ofrecognition and processing of substrates other than phenol.The biphasic nature of phenol disappearance, as moni-

tored by HPLC, suggests that accumulation of the endproduct might inactivate the enzyme, and therefore thepresence of the downstream enzyme, catechol 1,2 dioxy-genase (C1,2O) is necessary to shift the equilibrium towardsthe detoxification reactions. Product formation couldtherefore be assayed indirectly, by monitoring the activityof both enzymes. Preliminary results indicate that themultienzyme complex can efficiently attack methylated aswell as chloro-phenols and that a specific hydroxylationoccurs at the C2 position of substituted phenols. Furtherinvestigations are being pursued in order to confirm suchobservations.

Acknowledgements

The authors wish to thank Dr A. Peraino for HPLC experiments,

Dr T. Cacciatori for precious technical support, Dr Cavazzini for CD

results supervision, Dr M. G. Giuffrida and Dr A. Conti for amino

acid sequencing and ProfG. L. Rossi for helpful advice and discussion.

This project was supported by the EC Biotechnology Programme

(CT960413), by grants from the MURST (60%) and from Consorzio

Interuniversiario per le Biotecnologie (CIB).

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the oxygenase component of phenol hydroxylase from acinetobacter radioresistens s13

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