Gene cloning and characterization of dihydrolipoamide dehydrogenase from Microbacterium luteolum: A useful enzymatic regeneration system of NAD+ from NADH

Journal of Bioscience and Bioengineering VOL. 109 No. 3, 218 – 223, 2010 www.elsevier.com/locate/jbiosc

Gene cloning and characterization of dihydrolipoamide dehydrogenase from Microbacterium luteolum: A useful enzymatic regeneration system of NAD + from NADH Junji Kurokawa, Manabu Asano, Shunsuke Nomoto, Yoshihide Makino, and Nobuya Itoh⁎ Department of Biotechnology, Faculty of Engineering (Biotechnology Research Center), Toyama Prefectural University, 5180 Kurokawa, Imizu, Toyama 939-0398, Japan Received 28 April 2009; accepted 9 September 2009

Dihydrolipoamide dehydrogenase (LPD), a useful biocatalyst for regenerating NAD+, was purified from Microbacterium luteolum JCM 9174, and the gene encoding LPD was cloned from the genomic DNA. The gene contained an opening reading frame consisting of 1395 nucleotides encoding 465 amino acid residues with a predicted molecular weight of 49912.1 Da, which displayed 36–78% homology to known LPDs. Moreover, the FAD- and NAD+-binding sites and the two catalytic residues in the LPDs were conserved. The enzyme was expressed in recombinant Escherichia coli cells and purified to homogeneity by column chromatography. LPD of M. luteolum (MluLPD) accepted not only lipoamide but also some artificial electron acceptors such as dichlorophenolindophenol (DCIP) and nitrotetrazolium blue (NTB), that is, it functions as a diaphorase. NAD+ demonstrated a strong activating effect on MluLPD, and the activity was 5.2 times higher than that without NAD+. The enzyme was suitable for regenerating NAD+ in biocatalytic reactions because of its high affinity for NADH (6.1 μM). An NAD+-regenerating system with MluLPD and laccase using 2,5-dimethoxy-1,4-benzoquinone as a hydrogen acceptor was demonstrated. © 2009, The Society for Biotechnology, Japan. All rights reserved. [Key words: Dihydrolipoamide dehydrogenase; Diaphorase; Microbacterium luteolum; NAD+-regeneration; Artificial electron acceptor; 2,5Dimethoxy-1,4-benzoquinone; Laccase; Bilirubin oxidase]

Dihydrolipoamide dehydrogenase (LPD) (EC 1.8.1.4) catalyzes the NAD+-dependent dehydrogenation of dihydrolipoamide and is composed of two identical subunits, which generally contain 1 mol of FAD as a prosthetic group and one redox-active disulfide per subunit (1, 2). LPDs are generally sensitive to divalent cations such as Hg2+ and Cu2+ due to the presence of cysteine residues at the active site. The enzyme belongs to the flavin-containing pyridine nucleotidedisulfide oxidoreductase family, which includes glutathione reductase (EC 1.8.1.7) (1, 3), thioredoxin reductase (EC 1.8.1.9) (1, 4), and mercury (II) reductase (EC 1.16.1.1) (5). The enzyme is known to be an integral component of four multi-enzyme complexes: pyruvate dehydrogenase, 2-oxoglutarate dehydrogenase, branched-chain 2oxoacid dehydrogenase, and glycine decarboxylase (6–8). LPD catalyzes the final step of the sequential dehydrogenation of 2-oxo acids with regeneration of the lipoyl group from a dihydrolipoyl group using NAD+. LPDs react with not only lipoamide using NAD(H) as a physiological coenzyme but also many artificial electron acceptors including dichlorophenol indophenol (DCPIP), menadione, nitrotetrazolium blue (NTB), and ferrocene, functioning as a diaphorase (EC 1.6.99.-),

⁎ Corresponding author. Tel.: +81 766 56 7500x560; fax: +81 766 56 2498. E-mail address: [email protected] (N. Itoh).

which are used as a diagnostic reagent for colorimetric determination of NAD(P)H as well as many dehydrogenases (9, 10). This paper describes the properties of Microbacterium luteorum lipoamide dehydrogenase (MluLPD), which was cloned from M. luteolum and expressed in Escherichia coli, and its possible application to biocatalytic reactions not only to detect dehydrogenase activity but also to regenerate NAD+ using artificial electron acceptors by a coupled reaction with laccase (EC 1.10.3.2). MATERIALS AND METHODS Bacterial strains, plasmids, and culture conditions M. luteolum stain JCM 9174 was grown aerobically at 30 °C in medium containing 1.5% (w/v) peptone, 0.5% yeast extract, 0.5% NaCl, 0.3% sodium glutamate, and 1% sucrose (pH 7.0). E. coli JM109 was used as a host strain for cloning and expression of the mlulpd gene of M. luteolum. E. coli cells were cultivated at 37 °C in Luria–Bertani (LB) medium (1% tryptone, 0.5% yeast extract, and 0.5% NaCl, pH 7.0) containing 0.1 mg/ml ampicillin (total volume: 200 ml). For induction of the gene under the control of the lac promoter, 1 mM isopropyl-β-thiogalactopyranoside (ITPG) was added to LB medium. Enzyme assay LPD activity was assayed spectrophotometrically at 25 °C by measuring the decrease in the absorbance of NADH at 340 nm in a reaction mixture consisting of 2.0 mM lipoamide, 0.2 mM NADH, and 0.2 mM NAD+ (activator for LPD) in 50 mM potassium phosphate buffer (KPB) (pH 6.0). An extinction coefficient of 6.22 mM− 1 cm− 1 was used for NADH in the calculation of activity. Diaphorase activity was assayed at 25 °C in 50 mM KPB (pH 6.0) containing 0.2 mM NADH and 0.1 mM substrate [dichlorophenol indophenol (DCPIP), potassium ferricyanide, methylene blue, iodonitrotetrazolium chloride (INT), 1-methoxy phenazine methosulfate (PMS),

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DIHYDROLIPOAMIDE DEHYDROGENASE FOR NAD+ REGENERATION

resorufin, menadione, 2,5-dimethoxy-1,4-benzoquinone (2,5-DMBQ), 2,6-DMBQ) ] or 0.03 mM substrate [nitrotetrazolium blue (NTB), sulfonated tetrazolium (WST-1), resazurin)]. These compounds have the following absorption coefficients (nm, mM− 1 cm− 1): DCPIP (ɛ600, 16.1), potassium ferricyanide (ɛ420, 1), methylene blue (ɛ570, 13.1), INT (ɛ490, 15), 1-methoxy PMS (ɛ505, 2.84), NTB (ɛ530, 36), WST-1 (ɛ438, 37), resorufin (ɛ569, 8.25), and resazurin (ɛ600, 40.1). Diaphorase activity was calculated from the decrease/increase in absorption of the substrate at each wavelength. Menadione, 2,5-DMBQ, or 2,6-DMBQ reduction was assayed spectrophotometrically to observe the oxidation rate of NADH to NAD+. The reduction of ferrocene and ferrocenecarboxylic acid was assayed by measuring the amount of NADH and NAD+ by HPLC (11). One unit of enzyme was defined as the amount that converted 1 μmol of NADH or substrate in 1 min under these conditions. Purification of MluLPD from M. luteolum All purification procedures were performed at 4 °C in 20 mM KPB (pH 7.0) containing 1 mM 2-mercaptoethanol, 1 mM MgCl2, and 10% glycerol, unless otherwise indicated. The washed M. luteolum cells (15.8 g, wet weight) collected from 3 l of culture broth were suspended in 32 ml of buffer and then disrupted using an ultrasonic oscillator (INSONATOR 201 M; Kubota Corp., Tokyo, Japan). After centrifugation (13,000 × g , 30 min), the resulting supernatant was fractionated with solid ammonium sulfate. The precipitate obtained with 0 to 50% saturation of ammonium sulfate was collected, dialyzed against buffer (pH 7.0), and applied to a DEAE-Toyopearl 650 M (Tosoh Co., Ltd., Tokyo, Japan) column (22.5 × 625 mm) that had been equilibrated with the buffer. The enzyme was eluted using a linear 0 to 0.8 M NaCl gradient in the same buffer. The fractions with high enzyme activity were collected, and then the enzyme was loaded onto a Bioassist Q (Tosoh) column that had been equilibrated with buffer (pH 6.5) and was connected to an analytical high-performance liquid chromatography (HPLC) system. The enzyme was eluted with a linear 0 to 0.8 M NaCl gradient in the same buffer at a flow rate of 1.0 ml/min. The enzyme was again subjected to the same column using the buffer (pH 6.3), and eluted with a linear 0 to 0.6 M NaCl gradient in the same buffer at a flow rate of 0.7 ml/min. Next, the enzyme solution was applied to a TSK-gel G3000SW (Tosoh) gel filtration column (21.5 mm × 30 cm) that had been equilibrated with buffer (pH 7.0) and eluted at a flow rate of 1.5 ml/min. The fractions with high enzyme activity were collected, and then the enzyme was loaded onto a Bioassist Q column that had been equilibrated with buffer (pH 7.0), followed by elution with a linear 0 to 0.6 M NaCl gradient in the same buffer at a flow rate of 0.5 ml/min. The enzyme solution obtained was used as the partially purified enzyme for N-terminal and internal amino acid sequencing. Partial NH2-terminal and internal amino acid sequences of MluLPD The enzyme was electrophoresed on an SDS-PAGE gel, transferred to a polyvinylidene difluoride membrane (Bio-Rad) using a semidry electroblotting apparatus (NA-1512; Nippon Eido, Tokyo, Japan), and then stained with Coomassie brilliant blue G-250. The amino acid sequence at the N-terminal end of the enzyme on the polyvinylidene difluoride membrane was determined by using a HP G1005A protein sequencing system (Hewlett-Packard), and internal amino-acid sequences were determined by the APRO Life Science Institute (Tokushima, Japan). Cloning and DNA sequencing of the mlulpd gene M. luteolumJCM9174 genomic DNA was prepared as follows. The cells were cultured in 100 ml of LB medium, collected, and then disrupted using a Multi-Beads Shocker (MB-445U(S), Yasui Kikai, Osaka, Japan) with 500 μl of TE buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA) and 500 mg of glass beads (diameter: 0.5 mm) for 6 cycles of 30 s of 2,500 rpm agitation at 4 °C with 30 s intervals. After centrifugation, 500 μl of supernatant was mixed completely with 500 μl of TE-saturated phenol. The suspension was then centrifuged, and the aqueous phase was recovered. After treatment with RNase A (Nippon-Gene, Toyama, Japan), the DNA was precipitated using ethanol and resuspended in 100 μl of TE. Based on the N-terminal and internal amino acid sequences of the purified MluLPD (PHYDVVILGA, ADFFGISGEFTI), oligonucleotide primers of Mlu_lpdf (5′-CCNCAYTAYGAYGTNGTNATHYTNGGNGC-3′), and Mlu_lpdr (5′-AAYTCNCCNSWDATNCCRAARAARTCNGC-3′) were synthesized (Nippon EGT, Toyama, Japan). Polymerase chain reaction (PCR) was carried out using these primers and ExTaq DNA polymerase (Takara Bio Inc., Shiga, Japan) with M. luteolum genomic DNA as a template. The DNA fragments in the mixture were then cloned into a TOPO vector using a TOPO TA cloning kit (Invitrogen, CA, USA). Transformant colonies on the agar plate were subjected directly to PCR using the provided primers of M13 forward (− 20) and M13 reverse. Plasmids were prepared from the colonies amplifying an approximately 0.42-kbp fragment, which was expected from the homologous amino acid sequence of Mycoplasma bovis LPD. By sequencing analysis of the inserts of purified plasmids, the partial internal sequence of a probable LPD was found, according to homology with M. bovis LPD. Primers MluLPD-1R (5′-AGAAGTACTGGGGCGGTGTCTGCCTC-3′) and MluLPD-1F (5′-CTTCGATGATGGCCGTGGAGAGGCCG-3′), which corresponded to the partial internal sequences of the insert, were synthesized (Nippon EGT). Next, M. luteorum genomic DNA was partially digested using Sau3AI (Roche), size fractionated (3 to 7 kb) through a 1% agarose gel, and purified using a Wizard SV Gel and PCR Clean-Up System (Promega, WI, USA). The gene fragment mixture was then ligated into the BamHI restriction site of plasmid pUC118. E. coli JM109 cells were transformed with the ligation mixture, and a large number of the colonies on LB agar medium with 100 μg/ml ampicillin were collected. Plasmids were prepared from the genomic library transformants, and then inverse PCR was carried out on the plasmid mixture as the template using primers MluLPD-1R and MluLPD-1F, which had been phosphorylated

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using T4 polynucleotide kinase in advance (New England BioLabs, MA, USA). The amplified products were self-ligated and electroporated into E. coli JM109. The DNA sequences of the purified plasmids were analyzed and the possible 5′ and 3′ terminus sequences of the coding region of LPD were found by homology with the M. bovis LPD. The primers MluLPD-S1 (5′-CCCTCGTCGGAGCGATAGGCTTAACATATG-3′) and MluLPD-A1 (5′-CGCTTCGCTCGCTCAACCTGAAGCTTCGCG-3′), which were modified sequences adjacent to the coding region, were synthesized (note: the coding region of LPD was not modified by these primers). PCR were carried out using the high-fidelity polymerase KOD-plus (Toyobo, Osaka, Japan) and primers MluLPD-S1 and MluLPD-A1. To generate pMlulpd, the amplified DNA was digested with NdeI and HindIII, and cloned into the same sites of plasmid pUARH (12) to express the gene efficiently under the control of the lac promoter. The entire insert DNA sequences among the plasmids from 8 different colonies matched completely and were designated as pMlulpd. Expression and purification of MluLPD in E. coli E. coli JM109 cells harboring pMlulpd were grown at 37 °C in 1 l of LB medium with 50 μg/ml ampicillin and 1 mM IPTG for 12 h. Cells were harvested by centrifugation, suspended in 20 mM KPB (pH 7.0), and disrupted using an ultrasonic oscillator. Cell debris was removed by centrifugation at 14,000 × g , and the supernatant was filtered using a membrane filter (0.45 mm). The cell-free extract obtained was used for the further purification of recombinant MluLPD. The extract was first subjected to a DEAE-Toyopearl 650 M column (2.5 mm × 30 cm) chromatography, from which the enzyme was eluted with a linear gradient of 0 to 1 M NaCl in buffer (400 ml). The fractions exhibiting activity were mixed with ammonium sulfate up to a concentration of 1.2 M and applied to a ButylToyopearl 650 M (Tosoh) column (2.5 × 30 cm) that had been equilibrated with 1.2 M ammonium sulfate in buffer (pH 7.0). The enzyme was eluted with a linear 1.2 to 0 M ammonium sulfate gradient in buffer. The fractions with high enzyme activity were desalted using a Centriprep YM-30 filter unit (Millipore, molecular weight cut-off of 30,000 Da). The obtained enzyme solution was used as the purified enzyme for characterization. Molecular weight and SDS-PAGE The molecular weight of the enzyme was determined by analytical HPLC with a TSK-Gel G3000SWXL (Tosoh) column (7.8 mm × 30 cm) at a flow rate of 0.8 ml/min with 50 mM Tris–HCl (pH 7.0) containing 0.1 M NaCl. The molecular weight of the native enzyme was determined by comparison with the retention times of standard proteins. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was performed in a 12% polyacrylamide slab gel with a Tris-glycine buffer system as described by Laemmli (13). The molecular weight of the enzyme subunit was determined by comparison with the relative mobility of standard proteins. Determination of flavin The purified MluLPD was treated with 50% acetonitrile after boiling for 5 min according to the method of Youn et al. (14) followed by centrifugation. The yellowish supernatant was used to characterize and determine the content of flavin by HPLC using authentic compounds with a TSK GEL ODS-80Ts column (Tosoh). HPLC analysis was performed under isocratic conditions with a mobile phase of acetonitrile/water/diluted trifluoroacetic acid (10% v/v)/phosphoric acid (14:84:1.5:0.09) at a flow rate of 1.0 ml/min. Authentic compounds were detected at 260 nm, and the retention times of riboflavin, FMN, and FAD were 9.9, 6.6, and 3.9 min, respectively. Reaction mixture to regenerate NAD+ from NADH The reaction mixture to generate NAD+ from NADH consisted of 0.2 M KPB (pH 6.0), 5.0 mM NADH, 0.01– 0.1 mM 2,5-DMBQ, 1.2 units of MluLPD (for lipoamide), and 100 units of laccase in a total volume of 1 ml, and the reaction was performed at 30 °C with mixing. The laccase activity was assayed spectrophotometrically at 30 °C by measuring the formation of quinonimine dye at 505 nm from phenol and 4-aminoantipyrine (pH 4.5) in accordance with the manufacturer's protocol. In order to relate its activity with that of laccase, bilirubin oxidase (EC 1.3.3.5) activity was assayed under the same conditions above except KPB (pH 7.0) was used. One unit of the enzyme was defined as the amount that increased the absorbance at 505 nm by 0.1 optical density per minute. Oxidation of secondary alcohol coupled to NAD+ regenerating system To evaluate the efficiency of the NAD+-regeneration system, 2-octanol was oxidized into 2-octanone. The reaction mixture consisted of 20 mM KPB (pH 5.0–7.0), 10 mM (R, S)2-octanol, 1 mM NAD+, 0.1 mM 2,5-DMBQ, 5 units of Leifsonia alcohol dehydrogenase (LSADH), which catalyzes the stereospecific oxidation of (R)-2-octanol (15), 6 units Mlulpd and 100 units of laccase or 1.2 units of bilirubin oxidase in a total volume of 1 ml in a 2 ml-polypropylene tube. The reaction was conducted for 3 h at 30 °C with shaking (1200 rpm) in a Bioshaker M·BR-022UP (Taitec, Saitama, Japan). 2-Octanol and 2-octanone were quantitatively measured by gas chromatography (GC) using a Shimadzu GC-14 system equipped with a column (0.25 × 200 cm) packed with Thermon 1000 (5% on Chromosorb W) and a flame ionization detector. Nitrogen was used as the carrier gas at a flow rate of 50 ml/min, and the injection and detection temperatures were 220 °C. The column temperature was isothermally maintained at 80 °C. Under these conditions, 2-octanol and 2-octanone were detected at 3.4 and 7.1 min, respectively. Chemicals SDS-PAGE molecular weight standards (low range) were purchased from Nippon Bio-Rad Lab., Tokyo, Japan. The marker protein kit for HPLC was obtained from Oriental Yeast Co., Ltd., Tokyo. Laccase from Trametes sp. (Daiwa Y120, 120,000 U/g) was obtained from Amano Enzyme Co., Ltd., Nagoya, Japan, and bilirubin oxidase from Myrothecium verrucaria and tyrosinase (EC 1.14.18.1) from mushrooms were obtained from Sigma-Aldrich Japan. All other reagents were of analytical grade.

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Isolation of the mlulpd gene and construction of pMlulpd During the course of screening microorganisms that could convert some ketones to chiral alcohols, it was found that M. luteolum not only produced a target enzyme but also a detectable amount of diaphorase. General analysis of the enzyme indicated that the enzyme was an LPD and useful for oxidizing NADH to NAD+ using artificial electron acceptors. To apply MluLPD in an NAD+-regenerating system, it was first necessary to isolate the mlulpd gene and efficiently express it in E. coli cells. Although the direct purification of the enzyme form M. luteolum cells was incomplete and the final preparation of the enzyme contained contaminating proteins (data not shown), the MluLPD protein in a SDS-PAGE gel was transferred onto a PVDF membrane and then subjected to protein sequencing. The N-terminal amino acid sequence of MluLPD was PHYDVVILGA, and one of the internal amino acid sequences of the enzyme after lysyl endopeptidase treatment was ADFFGISGEFTI, which showed high homology with other LPD proteins. To isolate the gene from M. luteolum, a 0.42kb fragment containing an approximately 0.2 kb fragment of the mlulpd gene was obtained by PCR amplification using degenerate oligonucleotide primers designed from the N-terminal and internal amino acid sequences. The additional 5′ and 3′ terminal sequences of the coding region of mlulpd were amplified by inverse PCR. Sequence analysis indicated that the initiation codon (ATG) was preceded by a putative Shine-Dalgarno (SD) sequence (GATAGG). The open reading frame of mlulpd consisted of 1395 nucleotides encoding a protein of 465 amino acid residues with a predicted molecular

weight of 49912.14 Da. The deduced N-terminal (PHYDVVILGA) and internal amino acid sequences (ADFFGISGEFTI) of the enzyme were in agreement with the sequences that were used to prepare the PCR primers. The purified protein did not contain methionine at the N-terminus, indicating that the protein may be subjected to a post-translational modification. The nucleotide sequence reported in this paper is available from the DDBJ, EMBL, and GenBank databases under the accession number AB447982. A search of protein sequence databases (DDBJ, GenBank, EMBL, and PIR) revealed that MluLPD had a high homology with several LPDs of prokaryotes and eukaryotes, namely, 78.2% (identity) with Frankia sp. (EMBL accession number: ZP_00571989), 76.6% with Mycobacterium vanbaalenii (ZP_01204174), 73.5% with Rhodococcus sp. (YP_702105), 72.3% with Nocardia farcinica (YP_121481), 36.8% with pig (NP_999227), and 36.4% with human (P09622). The amino acid sequence alignment with other LPDs is shown in Fig. 1. FAD and pyridine nucleotide binding motifs (GXGXXG) were located at the Nterminus and roughly in the middle of the polypeptide chain, respectively. Two catalytic cysteine residues (GGXCXXXGCXP) involved in the redox-active disulfide site near the N-terminus and HisGlu pair residues (HXXXXE) located near the C-terminus were fully conserved in the MluLPD sequence. These four motifs are essential for the flavin-containing pyridine nucleotide-disulfide oxidoreductase activity (14). Expression and purification of MluLPD For the production of MluLPD in E. coli, the MluLPD-encoding region amplified by PCR was subcloned into pUARH (12) to generate pMlulpd. The pMlulpd

FIG. 1. Alignment of LPDs of M. luteolum (MluLPD, DDBJ: AB447982), Frankia sp. (EMBL: ZP_00571989), Mycobacterium vanbaalenii (ZP_01204174), Rhodococcus sp. (YP_702105), Nocardia farcinica (YP_121481), pig (NP_999227), and human (P09622). FAD and pyridine nucleotide binding motifs (GXGXXG) are underlined. Two catalytic cysteine residues (GGXCXXXGCXP) and His-Glu pair residues (HXXXXE) are indicated by a dashed underline and asterisk, respectively. Identical and similar residues are shown by white letters on black and grey backgrounds, respectively.

DIHYDROLIPOAMIDE DEHYDROGENASE FOR NAD+ REGENERATION

VOL. 109, 2010 TABLE 1. Summary of the purification of recombinant MluLPD from E. coli.

Crude extract DEAE-toyopearl 650M Butyl-toyopearl 650M

Total activity (U)

Total protein (mg)

Specific activity (U/mg)

Recovery of activity (%)

Purification (fold)

259.9 101.7 85.2

105.0 4.8 1.4

2.5 21.2 59.2

100.0 39.1 32.8

1.0 8.6 23.9

plasmid contained the mlulpd gene in the proper direction behind the lac promoter, which conferred high activity in the presence of IPTG. A sequence (GGAGAG) corresponding to the consensus sequence for the ribosome binding site for the E. coli translation system was located 7 bp upstream from the ATG codon of mlulpd. This system succeeded in producing a high expression level of the mlulpd gene in E. coli JM109 cells, and the content of MluLPD reached approximately 4% of the soluble proteins of E. coli (Table 1). The enzyme was easily purified from the recombinant E. coli cells using ion exchange and hydrophobic interaction chromatography with a yield of 32.8% (Table 1). The purified fractions showed a strong yellow color, suggesting that MluLPD could be a flavin-containing protein. Determination of flavin The absorption spectrum of MluLPD was measured by scanning the absorbance between 250 and 600 nm. The enzyme showed characteristic peaks at 270, 350, and 457 nm (data not shown), which corresponded to typical spectra of flavincontaining proteins. The flavin component was extracted from MluLPD and analyzed by HPLC, which indicated that the flavin component of MluLPD was FAD, and the content was calculated to be 1.34 mol per 1 mol of subunit of the enzyme. Enzyme properties The apparent molecular weight of recombinant MluLPD was determined by gel filtration using a TSK-Gel G3000SW column as described in Materials and methods. It was concluded that the molecular weight of MluLPD in its native form was approximately 102 kDa. The recombinant protein showed a single band following SDS-PAGE and the molecular weight of the protein was approximately 50 kDa. Comparison of this value with the theoretical molecular weight suggests that MluLPD exists as a dimeric protein in its native state. The optimal pH for MluLPD was found to be around 6.0, and it showed high activity in a pH range of 6.0 to 7.0 (Fig. 2). The purified enzyme showed activity toward not only lipoamide but also some artificial electron acceptors including dichlorophenol indophenol (DCPIP), ferrocene, ferrocene carboxylic acid, sulfonated tetrazolium (WST-1), iodonitrotetrazolium chloride (INT), nitrotetrazolium blue (NTB), 1-methoxy phenazine methosulfate (PMS),

FIG. 2. Optimal pH of MluLPD. Measurements were done in 100 mM buffer containing 2 mM lipoamide and 0.2 mM NAD+ and NADH. Buffers used were citrate-K2HPO4 (pH 4.5–6.0), KPB (pH 5.5–8.0), and Tris–HCl (pH 8.0 and 8.5).

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TABLE 2. Substrate specificity of MluLPD. Substrate Lipoamide DCPIP Ferrocene Ferrocenecarboxylic acid Potassium ferricyanide Methylene blue WST-1 INT NTB 1-Methoxy PMS Resazurin Resorufin Menadione 2,5-DMBQ 2,6-DMBQ

Relative activity (%) 100 233.4 3.6 4.1 ND ND 39.2 19.3 2.9 248.1 155.8 19.1 2.9 4.2 8.0

ND: not detected.

resazurin, resorufin, menadione, 2,5-dimethoxy-1,4-benzoquinone (2,5-DMBQ), and 2,6-DMBQ, but not toward potassium ferricyanide or methylene blue (Table 2). The Vmax and Km values for lipoamide in the presence of 0.2 mM NAD+ were calculated from a Lineweaver–Burk plot to be 120.5 μmol/ (min∙mg) and 3.3 mM, respectively; the Km value for NADH was 6.1 μM, which is the lowest value ever reported for LPDs. The enzyme showed no activity toward NADPH. MluLPD was thermostable, and the remaining activity after thermal treatment at pH 7.0 for 15 min was 90% at 55 °C, 80% at 60 °C, 70% at 65 °C, and 45% at 70 °C. Activation of Mlulpd by NAD+ As shown in Fig. 3, MluLPD exhibited 5.2 times higher activity in the presence of 0.2 mM NAD+ than in the absence of NAD+ for lipoamide reduction. Activation by NAD+ depending on sources is a well-known phenomenon for LPDs (14, 16), although the saturation curve in the absence of NAD+ was similar to those of allosteric enzymes. The role of NAD+ in the activation of lipoamide reduction with NADH is discussed later. However, this activation was not observed for the diaphorase activity when artificial electron acceptors were used as a substrate. Regeneration of NAD+ by MluLPD, laccase, and electron acceptors The characteristics of MluLPD including its high thermal stability, wide range of optimal pH, activity toward artificial electron acceptors, and especially low Km value for NADH (6.1 μM), were suitable properties to regenerate NAD+ in an NAD+/NADH recycling system. To clarify this point, a coupling system with laccase using 2,5-

FIG. 3. NAD+ activation of Mlulpd in lipoamide reducing reaction. Lipoamide reduction was measured with 0.2 mM NAD+ (square) and without NAD+ (circle). Bars indicate standard deviation for three measurements.

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FIG. 4. Coupling reaction of MluLPD and laccase using 2,5-DMBQ as hydrogen acceptor for NAD+ regeneration.

DMBQ as an electron acceptor was developed (Fig. 4). Guillen et al. reported in detail the oxidation mechanism of methoxyhydroquinones by Pleurotus eryngii laccase (16), in which the enzyme catalyzes the conversion of hydroquinones to semiquinones and to quinones during oxidation. When MluLPD (diaphorase) was added to the laccase-oxidation process of methoxyhydroquinones, it was expected to regenerate NAD+. In fact, this system using 0.1 mM 2,5-DMBQ rapidly oxidized 5 mM of NADH to NAD+, and no detectable NADH remained in the reaction mixture after a 40-min reaction time (Fig. 5). In addition, it was found that the 2,5-DMBQ concentration in the reaction mixture (0.01 and 0.1 mM) apparently affected the rate of NADH oxidation. Since the upper limit of 2,5-DMBQ solubility in water was around 0.1 mM, this concentration was used in subsequent experiments. Oxidation of 2-octanol by LSADH coupled to NAD+-regenerating system In order to optimize the reaction conditions and to evaluate the NAD+-regenerating system with MluLPD and laccase, racemic 2octanol oxidation by LSADH was coupled with this system. LSADH is a suitable enzyme for this test since it can catalyze the oxidation of (R)form chiral alcohols over a wide range of pH (15). When the reaction pH was varied from 5 to 7, the best result was obtained at pH 7.0. However, the optimum pH of Trametes laccase was around 4.5, and the activity at pH 7.0 was very low. Therefore, 100 additional units of

FIG. 5. Regeneration of NAD+ from NADH by MluLPD and laccase coupling reaction with 0.01 mM (square) and 0.1 mM (circle) 2,5-dimethoxy-1,4-benzoquinone (2,5-DMBQ) as hydrogen acceptor. Amounts of NAD+ and NADH were measured by HPLC. Closed circle or square indicates NAD+ and open circle or square indicates NADH concentration, respectively.

FIG. 6. Oxidation of racemic 2-octanol by LSADH coupled to NAD+-regenerating system with MluLPD and laccase. Circle and square indicate 2-octanol and 2-octanone concentration, respectively. When laccase (100 units) was replaced with bilirubin oxidase (1.2 units), nearly identical results were obtained.

laccase were needed for the reaction. An alternative enzyme, bilirubin oxidase from M. verrucaria that has an optimum pH at 7, was also used in this system. In this case, 1.2 units of bilirubin oxidase were sufficient to obtain the same reaction rate, suggesting that the enzyme possessed the same enzymatic function as laccase. On the other hand, mushroom tyrosinase was inert in this system. As shown in Fig. 6, half of the 2octanol was converted to 2-octanone after a reaction time of 3 h via this process. DISCUSSION In general, LPD activity is inhibited by the substrate NADH and activated by NAD+ in the lipoamide reduction. Wilkinson and Williams studied this phenomenon in detail using E. coli LPD (17) and concluded that the substrate inhibition of NADH was consistent with its reduction of the active two-electron reduced enzyme intermediate to the inactive four-electron reduced form (dead-end complex). NAD+ is able to overcome this inhibition by reversal of this reduction and binding to prevent the enzyme from forming a deadend complex. The inhibition of MluLPD was also prevented by the addition of NAD+ to the reaction mixture (Fig. 3). Diaphorases, including LPD, are a ubiquitous class of flavincontaining enzymes that exhibit NAD(P)H dehydrogenase activity by concomitantly catalyzing the reduction of oxidized compounds including synthetic dyes, quinones, oxygen, and lipoamide. On the basis of the original functions of the enzymes, they are classified into several categories: NAD(P)H dehydrogenase (quinone) (EC 1.6.5.2), NADPH dehydrogenase (EC 1.6.99.1), NADH dehydrogenase (EC 1.6.99.3), or dihydrolipoamide dehydrogenase (EC 1.8.1.4). Diaphorase has been used for the colorimetric determination of NAD(P)H via its dehydrogenase activity in vivo or in vitro coupled with various dyes that act as hydrogen acceptors for NAD(P)H. Clostridium kluyveri is a bacterial source of diaphorase that has been used commercially. Recently, Chakraborty et al. (9, 10) confirmed that C. kluyveri produces not only dihydrolipoamide dehydrogenase (bfmBC) but also a small diaphorase (24,981 Da) (diaA); therefore, commercially available diaphorase from C. kluyveri may be a mixture of two different NAD(P) H dehydrogenases. From the viewpoint of NAD(P)+ regeneration in biocatalysis, NAD(P)H dehydrogenases are useful enzymes to regenerate NAD(P)+ by coupling with dyes/O2 or O2 (Fig. 7). Hirano et al. (18) found a thermostable NADH dehydrogenase (NADH oxidase) in Brevibacterium sp., and applied it to the resolution of racemic mandelic acid

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FIG. 7. Schematic reaction process for production of ketones/aldehydes from alcohols by coupled enzymatic redox reactions.

coupled with an (R)-specific NAD+-dependent mandelate dehydrogenase to obtain (S)-mandelic acid (99% ee). Equilibria in the NAD+dependent dehydrogenation of alcohols generally favor alcohol formation from ketones/aldehydes (19). Therefore, an efficient regeneration system for NAD+ is indispensable to use such enzymes in organic synthesis. As shown in Fig. 5, the MluLPD and laccase/quinone system was very effective at regenerating NAD+ from NADH. The properties of MluLPD, especially its low Km value towards NADH (6.1 μM) and activity with 2,5-DMBQ, were considered to be useful for maintaining the ratio NAD+/NADH in the reaction mixture at the maximum level for NAD+ regeneration. As shown in Fig. 6, we confirmed that this redox coupling system is applicable to control the equilibrium in NAD+-dependent dehydrogenation of alcohols for organic synthesis, although further optimization is necessary. References 1. Williams, C. H.: Flavin-containing dehydrogenases, pp. 106-129, in: P. D. Boyer (Ed.), The Enzymes, vol. 13, Academic Press, New York, 1976. 2. Carothers, D. J., Pons, G., and Patel, M. S.: Dihydrolipoamide dehydrogenase: functional similarities and divergent evolution of the pyridine nucleotide-disulfide oxidoreductases, Arch. Biochem. Biophys., 268, 409–425 (1989). 3. Willmore, W. G. and Storey, K. B.: Purification and properties of glutathione reductase from liver of the anoxia-tolerant turtle, Trachemys csripta elegans, Mol. Cell. Biochem., 297, 139–149 (2007). 4. Jeon, S. J. and Ishikawa, K.: Identification and characterization of thioredoxin and thioredoxin reductase from Aeropyrum pernix K1, Eur. J. Biochem., 269, 5423–5430 (2002). 5. Ghosh, S., Sadhukhan, P. C., Chaudhuri, J., Ghosh, D. K., and Mandal, A.: Purification and properties of mercuric reductase from Azotobacter chroococcum, J. Appl. Microbiol., 86, 7–12 (1999). 6. Mattevi, A., de Kork, A., and Perham, R. N.: The pyruvate dehydrogenase multienzyme complex, Curr. Opin. Struc. Biol., 2, 877–887 (1992).

7. Perham, R. N.: Domains, motifs, and linkers in the design of a multifunctional protein, Biochemistry, 30, 8501–8512 (1991). 8. Walker, J. L. and de Kork, A.: Glycine decarboxylase multienzyme complex. Purification and partial characterization from pea leaf mitochondria, J. Biol. Chem., 261, 2214–2221 (1986). 9. Chakraborty, S., Sakka, M., Kimura, T., and Sakka, K.: Cloning and expression of a Clostridium kluyveri gene responsible for diaphorase activity, Biosci. Biotechnol. Biochem., 72, 735–741 (2008). 10. Chakraborty, S., Sakka, M., Kimura, T., and Sakka, K.: Characterization of a dihydrolipoyl dehydrogenase having diaphorase activity of Clostridium kluyveri, Biosci. Biotechnol. Biochem., 72, 982–988 (2008). 11. Itoh, N., Matsuda, M., Mabuchi, M., Dairi, T., and Wang, J. C.: Chiral alcohol production by NADH-dependent phenylacetaldehyde reductase coupled with in situ regeneration of NADH, Eur. J. Biochem., 269, 2394–2402 (2002). 12. Makino, Y., Dairi, T., and Itoh, N.: Engineering the phenylacetaldehyde reductase mutant for improved substrate conversion in the presence of concentrated 2propanol, Appl. Microbiol. Biotechnol., 77, 833–843 (2007). 13. Laemmli, U. K.: Cleavage of structural proteins during the assembly of the head of bacteriophage T4, Nature, 227, 680–685 (1970). 14. Youn, H., Kwak, J., Youn, H. D., Hah, Y., and Kang, S.: Lipoamide dehydrogenase from Streptomyces seoulensis: biochemical and genetic properties, Biochim. Biophys. Acta, 1388, 405–418 (1998). 15. Inoue, K., Makino, Y., and Itoh, N.: Purification and characterization of a novel alcohol dehydrogenase from Leifsonia sp. strain S749: a promising biocatalyst for an asymmetric hydrogen transfer bioreduction, Appl. Environ. Microb., 71, 3633–3641 (2005). 16. Guillén, F., Muńoz, C., Gómez-Toribio, V., Martínez, A. T., and Martínez, M. J.: Oxygen activation during oxidation of methoxyhydroquinones by laccase from Pleurotus eryngii, Appl. Environ. Microbiol., 66, 170–175 (2000). 17. Wilkinson, K. D. and Williams, Jr., C. H.: NADH inhibition and NAD activation of Escherichia coli lipoamide dehydrogenase catalyzing the NADH-lipoamide reaction, J. Biol. Chem., 256, 2307–2314 (1981). 18. Hirano, J., Miyamoto, K., and Ohta, H.: Purification and characterization of thermostable H2O2-forming NADH oxidase from 2-phenylethanol-assimilating Brevibacterium sp. KU1309, Appl. Microbiol. Biotechnol., 80, 71–78 (2008). 19. Kroutil, W., Mang, H., Edegger, K., and Faber, K.: Biocatalytic oxidation of primary and secondary alcohols, Adv. Synth. Catal., 346, 125–142 (2004).