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Applied and Environmental Microbiology, May 2009, p. 3106-3114, Vol. 75, No. 10
0099-2240/09/$08.00+0     doi:10.1128/AEM.02707-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Cloning, Expression, Characterization, and Biocatalytic Investigation of the 4-Hydroxyacetophenone Monooxygenase from Pseudomonas putida JD1{triangledown} ,{dagger}

Jessica Rehdorf, Christian L. Zimmer, and Uwe T. Bornscheuer*

Department of Biotechnology and Enzyme Catalysis, Institute of Biochemistry, Greifswald University, Felix-Hausdorff-Strasse 4, D-17487 Greifswald, Germany

Received 26 November 2008/ Accepted 20 February 2009


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ABSTRACT
 
While the number of available recombinant Baeyer-Villiger monooxygenases (BVMOs) has grown significantly over the last few years, there is still the demand for other BVMOs to expand the biocatalytic diversity. Most BVMOs that have been described are dedicated to convert efficiently cyclohexanone and related cyclic aliphatic ketones. To cover a broader range of substrate types and enantio- and/or regioselectivities, new BVMOs have to be discovered. The gene encoding a BVMO identified in Pseudomonas putida JD1 converting aromatic ketones (HAPMO; 4-hydroxyacetophenone monooxygenase) was amplified from genomic DNA using SiteFinding-PCR, cloned, and functionally expressed in Escherichia coli. Furthermore, four other open reading frames could be identified clustered around this HAPMO. It has been suggested that these proteins, including the HAPMO, might be involved in the degradation of 4-hydroxyacetophenone. Substrate specificity studies revealed that a large variety of other arylaliphatic ketones are also converted via Baeyer-Villiger oxidation into the corresponding esters, with preferences for para-substitutions at the aromatic ring. In addition, oxidation of aldehydes and some heteroaromatic compounds was observed. Cycloketones and open-chain ketones were not or poorly accepted, respectively. It was also found that this enzyme oxidizes aromatic ketones such as 3-phenyl-2-butanone with excellent enantioselectivity (E >>100).


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INTRODUCTION
 
Baeyer-Villiger monooxygenases (BVMOs; EC 1.14.13.x) belong to the class of oxidoreductases and convert aliphatic, cyclic, and/or aromatic ketones to esters or lactones, respectively, using molecular oxygen (29). Thus, they mimic the chemical Baeyer-Villiger oxidation, which is usually peracid catalyzed and was first described by Adolf Baeyer and Viktor Villiger in 1899 (2). All characterized BVMOs thus far are NAD(P)H dependent and require flavin adenine dinucleotide (FAD) or flavin mononucleotide (FMN) as prosthetic group, which is crucial for catalysis.

Today, BVMOs are increasingly recognized as valuable catalysts for stereospecific oxidation reactions. These enzymes display a remarkably broad acceptance profile for nonnatural substrates. Besides conversion of a wide range of aliphatic open-chain, cyclic, and aromatic ketones, they are also able to oxygenate sulfides (16), selenides (27), amines (33), phosphines, olefins (5), aldehydes, and borone- and iodide-containing compounds (Fig. 1) (7).


Figure 1
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  		FIG. 1. Range of Baeyer-Villiger oxidations catalyzed by BVMOs.

Therefore, recombinantly available BVMOs are powerful tools in organic chemistry and demonstrate a high potential as alternatives to existing chemical technologies, where some of these reactions are difficult to perform selectively using chemical catalysts.

Except for this promiscuity in reactivity, high enantioselectivities, as well as regio- and stereoselectivities, make them interesting for the pharmaceutical, food, and cosmetic industries, where enantiomerically pure compounds are valuable building blocks. In addition, renunciation of peracids when applying enzymatic driven Baeyer-Villiger oxidations turns them into an ecofriendly alternative and led to a considerable interest for biotransformations using BVMOs on an industrial scale (1, 8, 13-15) during the past decades.

Already in 1948 it was recognized that enzymes catalyzing the Baeyer-Villiger reaction exist in nature (39). This was concluded from the observation that a biological Baeyer-Villiger reaction occurred during the degradation of steroids by fungi. Still it took 20 years for the first BVMO to be isolated and characterized (10). Thus far, 22 BVMOs have been cloned, functionally expressed, and characterized. In Fig. 2 their genetic relationships are illustrated, and all BVMOs are sorted into different classes on the basis of their substrate specificity. Only two BVMOs, the 4-hydroxyacetophenone monooxygenase (HAPMO) from Pseudomonas fluorescens ACB (19) and phenylacetone monooxygenase (PAMO) from Thermobifida fusca (11), converting arylaliphatic and aromatic ketones were described. The latter is the only thermostable BVMO and served as a model to elucidate the enzymatic mechanism (28).


Figure 2
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  		FIG. 2. Phylogenetic relationships within BVMOs. The sequences of 22 enzymes with confirmed BVMO activity were aligned, and an unrooted phylogenetic tree was generated using CLUSTAL W (v.1.81). Cycloketone-converting BVMO (solid lines), open-chain ketone-converting BVMO (dashed lines), and arylketone-converting BVMO (dash/dot lines). NCBI accession numbers of protein sequences: CHMO Acinetobacter, CHMO Acinetobacter calcoaceticus NCIMB 9871 (BAA86293); CHMO Xanthobacter, BVMO Xanthobacter sp. strain ZL5 (CAD10801); CHMO Brachymonas, CHMO Brachymonas petroleovorans (AAR99068); CHMO1 Arthrobacter, CHMO1 Arthrobacter sp. strain BP2 (AAN37479); CHMO2 Arthrobacter, CHMO2 Arthrobacter sp. strain L661 (ABQ10653); CHMO1 Rhodococcus, CHMO1 Rhodococcus Phi1 (AAN37494); CHMO2 Rhodococcus, CHMO2 Rhodococcus Phi2 (AAN37491); CHMO1 Brevibacterium, CHMO1 Brevibacterium sp. strain HCU (AAG01289); CHMO2 Brevibacterium, CHMO2 Brevibacterium sp. strain HCU (AAG01290); CPMO Comamonas, cyclopentanone monooxygenase Comamonas sp. strain NCIMB 9872 (BAC22652); CPDMO Pseudomonas, cyclopentadecanone monooxygenase Pseudomonas sp. strain HI-70 (BAE93346); CDMO R. ruber, cyclododecane monooxygenase Rhodococcus ruber SCI (AAL14233); BVMO Mycobacterium tuberculosis Rv3083, BVMO M. tuberculosis H37Rv (gene Rv3083) (CAA16141); BVMO M. tuberculosis Rv3049c, BVMO M. tuberculosis H37Rv (gene Rv3049c) (CAA16134); BVMO M. tuberculosis Rv3854c, BVMO M. tuberculosis H37Rv (gene Rv3854c) (CAB06212); BVMO P. putida KT2440, BVMO P. putida KT2440 (AAN68413); BVMO P. fluorescens DSM50106: BVMO P. fluorescens DSM50106 (AAC36351); BVMO Pseudomonas veronii MEK700, BVMO P. veronii MEK700 (ABI15711); STMO Rhodococcus rhodochrous, steroid monooxygenase R. rhodochrous (BAA24454); PAMO T. fusca, phenylacetone monooxygenase T. fusca (Q47PU3); HAPMO P. fluorescens ACB, 4-hydroxyacetophenone monooxygenase from P. fluorescens ACB (AAK54073); HAPMO P. putida JD1, 4-hydroxyacetophenone monooxygenase from P. putida JD1 (FJ010625 [the present study]).

We report here the amplification, cloning, functional expression, and characterization of a HAPMO from Pseudomonas putida JD1 oxidizing a broad range of aromatic ketones and further substrates.


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MATERIALS AND METHODS
 
Chemicals.
All chemicals were of highest purity and purchased from Fluka (Buchs, Switzerland), Sigma-Aldrich (Taufkirchen, Germany), Roth GmbH (Karlsruhe, Germany), and Merck (Darmstadt, Germany) unless otherwise specified. Restriction enzymes were obtained from New England Biolabs (Beverly, MA), and NADPH was from Codexis (Jülich, Germany). DEAE-Sepharose FastFlow was from GE Healthcare (Uppsala, Sweden), and Reactive Red 120-Agarose was from Sigma-Aldrich. All β-hydroxyketones, as well as 4-hydroxy-4-phenyl-2-butanone, were synthesized according to the method of Kourouli et al. (23), and 3-phenyl-2-ketones and 1-phenylpropylacetate were synthesized according to the methods of Rubottom and Kim (35) and Krebsfänger et al. (24), respectively. Corresponding ester standards of β-hydroxyketones for gas chromatography-mass spectrometry (GC-MS) analysis were prepared enzymatically according to the method of Kirschner and Bornscheuer (22).

Bacterial strains, culture conditions, and plasmids.
P. putida JD1 was kindly provided by David J. Hopper (Institute of Biological Sciences, University of Wales, Aberystwyth, Ceredigion, United Kingdom). Escherichia coli DH5{alpha} [{Delta}lacU169({phi}80lacZ{Delta}M15) hsdR17 recA1 end A1 gyrA96 thi-1 relA1] was obtained from Clontech (Mountain View, CA). E. coli JM109 [recA1 endA1 gyrA96 thi-1 hsdR17 supE44 relA {Delta}(lac-proAB) F– (traD36 proAB+ lacIq lacZ{Delta}M15)] was obtained from New England Biolabs (Beverly, MA). E. coli BL21(DE3) [F– ompT hsdSB (rB– mB–) gal dcm rne131 (DE3)] and Rosetta [F– ompT hsdSB(rB– mB–) gal dcm lacY1 (DE3)/pRARE (Camr)] and the plasmid pET22b(+) were purchased from Novagen (Darmstadt, Germany). E. coli BL21-CodonPlus (DE3)-RP [F– ompT hsdS (rB– mB–) dcm+ Tetr gal {lambda}(DE3) endA Hte (argU proL; Cam) (argU ileY leuW; Strep/Spec)] was from Stratagene (La Jolla, CA), and E. coli BL21 C41(DE3) [F– ompT gal hsdSB (rB– mB–) dcm lon {lambda}DE3] was obtained from OverExpress. OneShot TOP10 cells [F– mcrA {Delta}(mrr-hsdRMS-mcrBC) {phi}80lacZ{Delta}M15 {Delta}lacX74 recA1 araD139 {Delta}(ara-leu)7697 galU galK rpsL (Strr) endA1 nupG] for blue-white screening were purchased from Invitrogen (Paisley, United Kingdom). E. coli strains were routinely cultured in LB medium and, when necessary, supplemented with ampicillin (100 µg/ml) or chloramphenicol (25 µg/ml). The Chaperone plasmid set containing the plasmids pG-KJE8, pGro7, pKJE7, pG-Tf2, and pTF16 was purchased from TaKaRa Bio, Inc. (Otsu, Japan).

Genetic methods and sequence analysis.
Total genomic DNA (gDNA) from P. putida JD1 was amplified by using a GenomiPhi v.2 DNA amplification kit from GE Healthcare. Plasmid isolations (Fermentas, St. Leon-Roth, Germany), PCR purification, and gel extraction (Qiagen, Hilden, Germany) were performed according the protocols of the manufacturers. Standard procedures such as DNA cloning and manipulations were performed as described by Sambrook and Russell (36). DNA sequencing was conducted by GATC (Konstanz, Germany), and analyses were carried out using the software package VectorNTI from Invitrogen.

16S rRNA gene sequencing.
The nearly full-length 16S rRNA gene of the P. putida JD1 strain was amplified by PCR using the eubacterial 16S rRNA gene primers 27f and 1492r as described by Lane (26). For further information, see the supplemental material.

Cloning of the HAPMO-encoding gene.
Degenerate primers were designed on the basis of three peptides (including the N-terminal sequence) obtained by Edman degradation of the HAPMO as described by Tanner and Hopper (38) to perform gradient PCR using gDNA and OptiTaq DNA polymerase (Roboklon, Berlin, Germany). The sequences of the primers were as follows: PpJD1-4HAPMO-FW1deg, ATGCGCACCTAYAAYACCAC; PpJD1-4HAPMO-RV1deg, GCGCCGGTGCCGATSACSGCSACGCG; PpJD1-4HAPMO-RV2deg, ATCCAGGTGCCRTCRTCGCGGATGATGC; and PpJD1-4HAPMO-Cterm1deg, TACATGCAGAACATRTTSGGGAACTGSGGSACGGTCATRC. The PCR conditions were as follows: 3 min at 95°C; followed by 45 cycles consisting of 95°C for 1 min, 50 to 65°C for 30 s, and 72°C for 1.5 min; and a final extension step of 7 min at 72°C. Due to the use of degenerate primers the annealing step was performed between 55 and 65°C. The sequence of the PCR product was blasted in nonredundant databases. The most similar hit was the HAPMO from P. fluorescens ACB. Thus, it was assumed that the amplified product is indeed part of the sequence of the HAPMO gene from P. putida JD1. Since the missing C-terminal end of the gene could not be amplified using inverse PCR (32), SiteFinding-PCR was performed as an alternative approach as described Tan et al. (37). The cycler conditions are given in Table S1 in the supplemental material. As counter primers, two gene-specific primers were designed: GSP1 (AAAAAGCTTTTGCCTCGCTCAG) and GSP2 (TTGGCAGCTGCTCAAGGTCG). The amplified product was subcloned directly in the pCRII-TOPO vector, and OneShot TOP10 cells were transformed with this construct. After sequencing, the HAPMO gene was reamplified with nested PCR from gDNA using PfuPlus DNA polymerase and primers incorporating restrictions sites (NdeI and BamHI [underlined in the sequences below]) for direct cloning into the expression vectors pET22b(+) and pJOE4072.6. The sequences of the primers were: FW1-HAPMO, TTCATCGGGCAGATGCACGG; FW2-HAPMO-NdeI, CATATGAGAACCTACAACACCACTTTGGC; RV1-HAPMO, TTGTAATTATTACCTGGCGGCCAG; and RV2-HAPMO-BamHI, GGATCCTCAGGAAAGGCAGTAGTCGG. The amplified product was subcloned into the pCRII-TOPO vector, digested with NdeI and BamHI (37°C, 3 h), and ligated into NdeI/BamHI-linearized pET22b(+) and pJOE4072.6, respectively. The plasmids were named pJOEPpJD1HAPMO and pET22b(+)PpJD1HAPMO and transformed into E. coli cells, followed by sequencing to confirm correct sequences.

Expression of 4-HAPMO in E. coli.
E. coli JM109 was transformed with pJOE PpJD1HAPMO, and all BL21(DE3) strains, JM109 (DE3), and E. coli Rosetta were transformed with pET22b(+)PpJD1HAPMO. In all cases, expression was evaluated in 30 ml of LB or 2xYT medium (2% tryptone, 1% NaCl, 1% yeast extract) containing 100 µg of ampicillin/ml. For the expression in E. coli Rosetta, 25 µg of chloramphenicol/ml was also added. Induction, protein expression, and sample preparation for sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) were performed according to the method of Rehdorf et al. (34).

Coexpression with molecular chaperones.
E. coli JM109 and BL21(DE3) cells were transformed with chaperone-encoding plasmids (pGro7, pKJE7, pTf16, pG-KJE8, and pTf16). Cells were grown in 20 ml of LB containing 34 µg of chloramphenicol (LBcap)/ml at 37°C, and competent cells were prepared (36). These cells were transformed with pJOE constructs (for JM109) or pET22b(+) constructs [for BL21(DE3)] and selected on LBcap+amp. Expression was performed at 30°C as described above using LBcap+amp containing L-arabinose (in case of pGro7, pKJE7, and pTf16) at 0.5 mg/ml or 5 ng of tetracycline/ml (in the case of pG-Tf2) or L-arabinose and tetracycline (in the case of pG-KJE8) in the concentrations given above.

Enzyme purification.
Overexpressed HAPMO was purified to homogeneity in two chromatographic steps. For this, HAPMO was expressed in E. coli Rosetta (induction with 0.1 mM IPTG [isopropyl-β-D-thiogalactopyranoside] at an optical density at 600 nm of 1) using 2xYT-medium at 20°C for 6 h in a volume of 500 ml. Cells (4 g [wet weight]) were then harvested, washed twice with sterile sodium phosphate buffer (pH 7.0, 20 mM), and disrupted with a precooled French pressure cell (Polytec) at 96.5 x 103 kPa. The lysate was centrifuged for 30 min at 10,000 x g and 4°C. The supernatant (30 ml) was taken as crude extract, and purification steps were performed as described by Tanner and Hopper (38). Fractions containing HAPMO activity were pooled and concentrated (Amicon/Millipore, Schwalbach, Germany). All purification steps were performed at room temperature.

Kinetic measurements.
HAPMO activity was determined spectrophotometrically by monitoring the decrease of NADPH at 370 nm ({varepsilon} = 1.96 mM/cm). Reaction mixtures (1 ml) contained 50 mM Tris-HCl buffer (pH 8.0), 312.5 µM NADPH, 28 mU of enzyme, and 10 µl of 0.1 mM 4-hydroxyacetophenone in dimethyl formamide. The reaction was started by adding NADPH to the mixture. One unit of HAPMO is defined as the amount of protein that oxidizes 1 µmol of NADPH per min. All kinetic measurements were performed at 30°C in air-saturated buffer. The kinetic parameters for all derivatives were determined by using at least six substrate concentrations ranging from 1 µM to 60 mM and a fixed concentration of 312.5 µM NADPH. The Km and Vmax for NADPH were determined by using a fixed concentration of 500 µM 4-hydroxyacetophenone and NADPH in a concentration range from 5 to 625 µM. All concentrations were measured in triplicate, and data were calculated according to the Hanes-Wolff equation.

Determination of protein concentration.
The protein concentration was determined by using a BCA assay protein quantification kit from Uptima (Montluçon, France) with bovine serum albumin as a standard in sodium phosphate buffer (pH 7.0, 20 mM).

Effect of pH on activity.
A total of 28 mU of purified enzyme was preincubated in acetate buffer (pH 4.0 to 5.5), phosphate buffer (pH 7.5 to 8.0), and Tris-HCl (pH 7.5 to 9.0) at 25°C for 30 min. The activity was measured spectrophotometrically as described above using 312.5 µM NADPH and 0.5 mM 4-hydroxyacetophenone.

Thermal stability.
The half-life at various temperatures was studied by preincubating protein samples (28 mU of purified enzyme) in Tris-HCl buffer (pH 8.0, 50 mM) at 25, 30, and 35°C. After definite time intervals (10, 20, 30, and 40 min for 35°C; 0.5, 1, 2, and 3 h for 30°C; and 1, 2, 5, and 24 h for 25°C), samples were taken and centrifuged at 4°C, and the residual activity was assayed as described above.

SDS-PAGE.
SDS-PAGE was carried out according to the method described by Laemmli (25).

Biocatalysis.
Substrate specificity of HAPMO was investigated by using crude cell extract. For all experiments, E. coli Rosetta pET22b(+)PpJD14HAPMO cells were used. The following substrates were investigated: 4-hydroxy-2-octanone, 4-hydroxy-2-nonanone, 4-hydroxy-2-decanone, and 4-hydroxy-2-undecanone; 3-phenyl-2-butanone; 3-phenyl-2-pentanone; and 4-hydroxy-4-phenyl-2-butanone. 4-Hydroxyacetophenone served as positive control.

Cells were grown in 2xYTcap+amp medium at 37°C to an optical density at 600 nm of 1 to 1.5. HAPMO expression was induced by the addition of IPTG to a final concentration of 0.1 mM. Expression was performed at 20°C for 6 h. Cells were harvested and washed twice with sterile 50 mM Tris-HCl buffer (pH 8.0) and finally resuspended in the same buffer. Cell disruption was performed using a precooled French press (96.5 x 103 kPa), and cell debris was removed by centrifugation (30 min, 4°C, 10,000 x g). Biocatalysis reactions were performed in flasks at a volume of 5 ml at 25°C. To 5 ml of cell lysate 7 µmol of substrate and 0.2 mmol of NADPH were added. After defined time intervals (1, 2, 4, 6, and 12 h) samples were obtained, extracted with ethyl acetate, and dried over anhydrous sodium sulfate. Samples were analyzed by GC and GC-MS.

GC and GC-MS analysis.
Achiral GC-MS analyses were carried out on a GCMS-QP 2010 apparatus (Shimadzu Europa GmbH, Duisburg, Germany) with a BPX5 column (5% phenyl-/95% methylpolysilphenylen/siloxan; SGE GmbH, Darmstadt, Germany). The injection and detection temperatures were set to 220 and 300°C, respectively. Chiral analytics were done on a heptakis-(2,6-di-O-methyl-3-O-pentyl)-β-cyclodextrin column (Hydrodex-β-3P, 25 m by 0.25 mm; Macherey-Nagel, Düren, Germany) in a Hewlett-Packard GC 5890 series II (Agilent, Waldbronn, Germany). Injection and detection temperatures were set to 220°C.

Accession numbers.
The nucleotide sequence of the amplified gene of 4-hydroxacetophenone monooxygenase from P. putida JD1 and the associated protein sequence, as well as a 16S rRNA gene sequence from P. putida JD1, have been deposited under GenBank accession numbers FJ010625 for the 4-hydroxyacetophenone monooxygenase and FJ010624 for the 16S rRNA gene.


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RESULTS AND DISCUSSION
 
Amplification and cloning of the HAPMO gene.
In 2000, three protein fragments, including the N-terminal sequence, of a Baeyer-Villiger monooxygenase from P. putida JD1 were determined by using Edman degradation, but the identification of the encoding gene and the recombinant expression were not reported. A BLAST search of these protein fragments revealed an identity of ~80% toward a BVMO from P. fluorescens ACB (GenBank accession no. AAK54073). On the basis of this sequence information, we then designed degenerate primers in order to perform PCR at different annealing temperatures. After sequencing of a 1.6-kb amplificate, identification of typical BVMO sequence motifs, and determination of the three known peptides, it was certain that the amplified sequence of 536 amino acids was indeed part of the HAPMO gene. Unfortunately, various PCR methods, including inverse PCR, failed to obtain the missing gene fragments, and the SiteFinding-PCR (see Fig. S1 in the supplemental material) was investigated (for a detailed method information, see the study by Tan et al. [37]). With this SiteFinding-PCR, a 10.2-kb fragment containing the whole HAPMO gene (1,920 bp) and several other open reading frames located up- and downstream could be identified. Analysis of these flanking genes suggests that HAPMO belongs together with four other putative enzymes (an esterase, a dioxygenase, a reductase, and a dehydrogenase) and a putative LysR regulator protein to an operon (Fig. 3). Since 4-hydroxyacetophenone can be considered the key substrate, the following biodegradation pathway in P. putida JD1 can be proposed (Fig. 4). First, 3-hydroxyacetophenone is converted into the corresponding ester by HAPMO, followed by the formation of a hydroquinone, which is further degraded via 3-hydroxymuconic semialdehyde and maleylacetate to β-ketoadipate, similar to the pathway described by Moonen et al. (30). Hence, the enzymatic Baeyer-Villiger oxidation is the key reaction to produce a substrate for the subsequent ring cleavage and further degradation. It has already been observed that in Acinetobacter sp. strain NCIMB 9871 and Comamonas sp. strain NCIMB 9872 the gene encoding the BVMO is clustered on the chromosome together with a hydrolase and a dehydrogenase (9, 17), which we also found for a BVMO from P. putida KT2440 (31).


Figure 3
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  		FIG. 3. Organization of the gene cluster of the 10.2-kb DNA fragment of P. putida JD1 amplified using SiteFinding-PCR. Dehydrogenase, putative 3-hydroxymuconic semialdehyde dehydrogenase; reductase, putative maleylacetate reductase; dioxygenase, putative hydroquinone dioxygenase; esterase, putative 4-hydroxyphenyl acetate hydrolase; 4-HAPMO, 4-hydroxyacetophenone monooxygenase; LysR, regulatory protein (LysR family).


Figure 4
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  		FIG. 4. Postulated degradation pathway of 4-hydroxyacetophenone catalyzed by the putative enzymes of the P. putida JD1 operon.

Protein expression and purification.
The expression of HAPMO was investigated using E. coli JM109 (with plasmid pJOEPpJD1HAPMO), various E. coli BL21(DE3) strains, and E. coli Rosetta [with plasmid pET22b(+)PpJD1HAPMO]. Different medium compositions and expression temperatures, as well as the addition of FMN, which is a precursor in the synthesis of FAD, were studied. Complex (LB medium supplemented with 2.5% [wt/vol] glucose and 2.5% [wt/vol] K2HPO4·3H2O) and 2xYT media were suggested to be optimal for these experiments. Adding FMN to the culture had no effect on HAPMO expression. The pET22b(+) vector yielded a higher amount of protein compared to the pJOE construct but, unfortunately, the majority of protein was produced as inclusion bodies. To overcome this problem, the expression temperature was lowered from 30 to 20°C. In addition, coexpression analysis using molecular chaperones was performed, but the expression of soluble protein was not improved (data not shown). A comparison of different E. coli strains [with the pET22b(+)PpJD1HAPMO construct] revealed that the Rosetta strain gave best results, since it supplies rare tRNAs, yielding the highest amounts of soluble HAPMO under optimized conditions at 20°C in complex medium after 6 h. The crude protein was purified to homogeneity in a two-step procedure using ion-exchange chromatography, followed by a 2'-ADP-based affinity chromatography with a recovery of 57% and a purification factor of 43 (Table 1 and Fig. 5).


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  		TABLE 1. Purification of recombinant P. putida JD1 HAPMO from E. coli


Figure 5
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  		FIG. 5. Purification steps of HAPMO. Lanes: 1, lysate (1:10); 2, pooled active fractions from ion-exchange chromatography; 3, low-range marker (range, 6.5 to 66 kDa) from NEB; and 4, active fraction from affinity chromatography.

pH optimum and thermal stability.
The maximum activity was measured at pH 8 in Tris-HCl buffer, while at a pH of <6 the activity is very low (see Fig. S2 in the supplemental material). The highest thermal stability was found at 30°C. After 30 min at 35°C HAPMO possesses only 47% residual activity, and at 40°C it possesses even less than 1% (see Fig. S3 in the supplemental material). At 25°C HAPMO showed the highest activity and was found to be stable for at least 20 h.

Substrate specificity and kinetic measurements.
In order to explore the substrate specificity, a wide range of potential substrates were investigated. It was found that HAPMO displays a broad substrate specificity among aromatic ketones (Table 2). Conversion of aromatic substrates was already reported for HAPMO from P. fluorescens ACB (19, 20). Open-chain and cyclic aliphatic ketones, which are good substrates for some known BVMOs (21, 34), were poorly accepted by HAPMO from P. putida JD1. Furthermore, the enzyme is strongly NADPH dependent. Oxidation of 4-hydroxyacetophenone does not occur with NADH as a cofactor. Substrates of HAPMO can be grouped into three classes: (i) aromatic, (ii) heteroaromatic, and (iii) aliphatic compounds (Table 2). The substances that did not show significant activity (<0.1 U/mg of protein) at a concentration of 5 mM were 2-acetylpyridine, 3-acetylpyridine, 4-acetylpyridine, 4-hydroxy-3-methoxyacetophenone, 1-indanone, cyclohexanone, cyclopentanone, acetylindole, progesterone, acetone, hydroxyacetone, dihydroxyacetone, (endo/exo)-acetylnorbornene, and 4-decanone.


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  		TABLE 2. Kinetic parameters of HAPMO from P. putida JD1a

Conversion of aromatic compounds.
HAPMO from P. putida JD1 preferentially oxidizes acetophenone derivatives. Like HAPMO from P. fluorescens ACB, the highest catalytic activity could be observed with compounds bearing an electron-donating group in the para position of the aromatic ring. This suggests that the functional position at the aromatic ring (para, meta, or ortho) is crucial for substrate recognition and affinity and influences Km and kcat values, which is demonstrated for the three hydroxyacetophenone compounds (Table 2). Still, substitutions on the phenyl ring and modifications at the aceto function were accepted by HAPMO. Conversion of 4-hydroxyacetophenone (Km = 47 µM) and acetophenone (Km = 384 µM) was already reported by Tanner and Hopper (38), and Km values correlate well with our data (Table 2). Also, compared to HAPMO from P. fluorescens ACB (18), most of the values were in the same range. Interestingly, 4-nitroacetophenone is not converted by P. fluorescens ACB HAPMO, whereas P. putida JD1 HAPMO oxidizes it with a Km of 313 µM. A nitro group reduces the electron density of the aromatic ring through the negative mesomeric and inductive effect (–M and –I), while methyl, hydroxy, and amino functions possess a positive inductive (+I) or mesomeric effect (+M). The maximal turnover number for aromatic compounds was reached with 2'-hydroxyacetophenone (21.2 s–1), followed by methyl-4-tolyl sulfide (20 s–1), acetophenone (16.3 s–1), and trifluoroacetophenone (15.4 s–1). Kinetic data found for other aromatic substrates were in the same range (1.0 to 11.3 s–1), which indicates that substitutions on the ring do not influence maximal turnover numbers significantly. Only in response to 3-methoxyacetophenone does HAPMO show a low activity. The lowest Km values could be observed with 4-hydroxyacetophenone (38.1 µM) and 4-aminoacetophenone (5.6 µM), whereas for more bulky and hydrophobic substituents (4-methyl- and 4-methoxyacetophenone) the Km values were much higher. The structure of the aceto function also influences the affinity and rate of catalysis: while benzaldehyde has a Km similar to that of acetophenone, the turnover number is much smaller. An additional methyl group (in propiophenone) increases the substrate affinity with a much lower Km, but kcat remains in the range of benzaldehyde. More bulky substituents such as butyrophenone lead to a drastic decrease in affinity, indicating that the propionyl residue might be optimal for catalysis.

HAPMO also catalyzes sulfoxidation.
In addition to oxidation of ketones and aldehydes, some BVMOs are also able to catalyze the sulfoxidation of a variety of sulfides (6), which can be interesting chiral building blocks for the pharmaceutical chemistry. Here, methyl-4-tolyl sulfide was tested as a substrate for HAPMO. This prochiral sulfide was oxidized with a kcat of 20 s–1, while the Km value was rather high (18.8 mM), which is in contrast to the affinity toward the corresponding ketone 4-methylacetophenone. This differs from the HAPMO from P. fluorescens ACB, which converts both methyl-4-tolyl sulfide and 4-methylacetophenone at similar Km and kcat values (20).

Conversion of heteroaromatic compounds.
Thus far, only a few BVMOs have been tested for conversion of heteroaromatic compounds, and until now only the HAPMO from P. fluorescens ACB has been shown to be active against them (18). Here, three pyridine substrates and one pyrrole substrate were investigated. While the 2-, 3-, and 4-acetylpyridines are not converted, 2-acetylpyrrole is oxidized to the ester. Obviously, the pyrrole ring is preferred over the pyridine. With a Km value of 103 µM, the affinity for 2-acetylpyrrole is even higher than for acetophenone and some derivatives.

Oxidation of aliphatic ketones.
Although aliphatic cyclic ketones and open-chain ketones are known to be good substrates for some BVMOs (4, 21, 34), HAPMO is able to convert only a few of them. HAPMO showed no activity at all against cyclohexanone, cyclopentanone, and 4-decanone, while 2-decanone and 2,4-pentanedione were oxidized with turnover rates in the same range as some aromatic substrates. 2-Decanone was converted with a fivefold lower Km value compared to 2,4-pentadione.

Biocatalysis.
For kinetic resolution of several racemic substrates, crude cell extract was used. Although the use of crude cell extract requires the addition of expensive NADPH as a cofactor, which makes this system problematic from an economic point of view, it assures that the substrates and products are not further metabolized by the whole cells. Biotransformations using resting cells—facilitating cofactor regeneration by glucose addition—were also performed but gave results similar to those obtained with crude extract. The compounds investigated were grouped into (i) aliphatic open-chain racemic ketones (4-hydroxy-2-octanone, 4-hydroxy-2-nonanone, 4-hydroxy-2-decanone, and 4-hydroxy-2-undecanone) and (ii) arylaliphatic racemic substrates (3-phenyl-2-butanone, 3-phenyl-2-pentanone and 4-hydroxy-4-phenyl-2-butanone). The results for conversion, enantiomeric excess, and enantioselectivity are summarized in Tables 3 and 4. As already mentioned, aliphatic open-chain ketones are oxidized poorly by HAPMO (Table 3). Both enantiomeric excess and conversion were very low for all four substrates. A slight increase of the product enantiomeric excess could be observed for 4-hydroxy-2-decanone (50% ee), while conversion was highest for 4-hydroxy-2-undecanone (8.6%). Due to the structure of 4-hydroxy-2-ketones, two different products are possible (acetate or methyl ester), but with this HAPMO only the acetate was formed. With respect to enantioselectivity, 3-phenyl-2-butanone was converted with an exceptionally high E value of >200 at 45.6% conversion, with an enantiomeric excess for the product, 1-phenylethylacetate, of 99.2% ee (Table 4). The conversion could be further increased to 66% (data not shown) by the addition of an ion exchanger (Lewatit MP64) at pH 8.5, which allows then for a dynamic kinetic resolution. However, 3-phenyl-2-pentanone and 4-hydroxy-4-phenyl-2-butanone were not converted at all. This narrow substrate spectrum and high enantioselectivity for one compound matches results reported for BVMOs from P. fluorescens DSM50106 (22) and P. putida KT2440 (12), but the HAPMO from P. putida JD1 is far more selective.


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  		TABLE 3. Biocatalysis of β-hydroxy-2-ketones using crude cell extract (E. coli Rosetta pET22b(+)PpJD1HAPMO)a


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  		TABLE 4. Biocatalysis of arylaliphatic ketones using crude cell extract [E. coli Rosetta pET22b(+)PpJD1HAPMO]a

In summary, we could successfully clone, express, and characterize a 4-hydroxyacetophenone monooxygenase from P. putida JD1. The key to successful identification of the encoding gene was the SiteFinding-PCR, which proved to be a powerful and reliable tool for amplifying the unknown flanking regions from gDNA. Besides the HAPMO from P. fluorescens ACB (which has only 83% sequence identity) and the PAMO from T. fusca, this HAPMO is actually the third described BVMO preferentially oxidizing arylaliphatic ketones. Interestingly, within this class, the HAPMO described here converts a broad range of para-substituted aromatic ketones bearing either electron-donating (OH, CH3, and NH2) or electron-withdrawing (NO2) groups with very low Km values and high kcat values. Furthermore, HAPMO converts 3-phenyl-2-butanone into 1-phenylethylacetate with excellent selectivity, making it a versatile enzyme for biocatalysis.


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ACKNOWLEDGMENTS
 
We thank the Deutsche Bundesstiftung Umwelt for a stipend to Jessica Rehdorf.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Biotechnology and Enzyme Catalysis, Institute of Biochemistry, Greifswald University, Felix-Hausdorff-Str. 4, D-17487 Greifswald, Germany. Phone: 49-3834-86-4367. Fax: 49-3834-86-80066. E-mail: uwe.bornscheuer{at}uni-greifswald.de Back

{triangledown} Published ahead of print on 27 February 2009. Back

{dagger} Supplemental material for this article may be found at http://aem.asm.org/. Back


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Applied and Environmental Microbiology, May 2009, p. 3106-3114, Vol. 75, No. 10
0099-2240/09/$08.00+0     doi:10.1128/AEM.02707-08
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