Applied Microbiology and Biotechnology

, Volume 87, Issue 1, pp 185–193

Asymmetric synthesis of (S)-3-chloro-1-phenyl-1-propanol using Saccharomyces cerevisiae reductase with high enantioselectivity

Authors

  • Yun Hee Choi
    • Division of BiotechnologyThe Catholic University of Korea
  • Hye Jeong Choi
    • Division of BiotechnologyThe Catholic University of Korea
  • Dooil Kim
    • Systems Microbiology Research CenterKorea Research Institute of Bioscience and Biotechnology
  • Ki-Nam Uhm
    • Equispharm Ltd.
    • Division of BiotechnologyThe Catholic University of Korea
Biotechnologically Relevant Enzymes and Proteins

DOI: 10.1007/s00253-010-2442-5

Cite this article as:
Choi, Y.H., Choi, H.J., Kim, D. et al. Appl Microbiol Biotechnol (2010) 87: 185. doi:10.1007/s00253-010-2442-5

Abstract

3-Chloro-1-phenyl-1-propanol is used as a chiral intermediate in the synthesis of antidepressant drugs. Various microbial reductases were expressed in Escherichia coli, and their activities toward 3-chloro-1-phenyl-1-propanone were evaluated. The yeast reductase YOL151W (GenBank locus tag) exhibited the highest level of activity and exclusively generated the (S)-alcohol. Recombinant YOL151W was purified by Ni-nitrilotriacetic acid (Ni-NTA) and desalting column chromatography. It displayed an optimal temperature and pH of 40°C and 7.5–8.0, respectively. The glucose dehydrogenase coupling reaction was introduced as an NADPH regeneration system. NaOH solution was occasionally added to maintain the reaction solution pH within the range of 7.0–7.5. By using this reaction system, the substrate (30 mM) could be completely converted to the (S)-alcohol product with an enantiomeric excess value of 100%. A homology model of YOL151W was constructed based on the structure of Sporobolomyces salmonicolor carbonyl reductase (Protein Data Bank ID: 1Y1P). A docking model of YOL151W with NADPH and 3-chloro-1-phenyl-1-propanone was then constructed, which showed that the cofactor and substrate bound tightly to the active site of the enzyme in the lowest free energy state and explained how the (S)-alcohol was produced exclusively in the reduction process.

Keywords

Antidepressant drugsChiral intermediateDocking modelEnantioselectivityReductase

Introduction

The asymmetric reduction of ketones to optically pure alcohols has attracted a great deal of attention due to their possible use as chemotherapeutic drugs and chiral building blocks. Many oxidoreductases (EC 1.1.1.-) from a variety of microorganisms may be capable of performing the reduction of carbonyl groups with chemo-, regio-, and stereoselectivity (Goldberg et al. 2007; Schroer et al. 2007). Enzymatic ketone reductions can be achieved using isolated enzymes (Goldberg et al. 2007; Inoue et al. 2005; Kaluzna et al. 2005) or whole-cell systems (Ema et al. 2006; Moore et al. 2007; Xu et al. 2006). Saccharomyces cerevisiae (baker's yeast) carbonyl reductases have been previously reported to exhibit catalytic activity, converting some ketone substrates into enantiomeric alcohols (Kayser et al. 2005; Kaluzna et al. 2005).

Biotransformation of ketone substrates with reductase enzymes requires the presence of the cofactor NAD(P)H. Cofactor regeneration can be conducted via an enzyme-coupled (Ema et al. 2006; Ernst et al. 2005; Moore et al. 2007) or a substrate-coupled (Inoue et al. 2005; Makino et al. 2005; Schroer et al. 2007) approach. The former approach is characterized by the use of a second enzyme that catalyzes the oxidation of a cosubstrate to regenerate the reduced cofactor, while the latter process employs only one enzyme for the production of the desired compound and the regeneration of the cofactor.

Fluoxetine belongs to a class of medications referred to as selective serotonin reuptake inhibitors, which have emerged as key medicines for the treatment of depression (Andrews et al. 2009; Vaswani et al. 2003). Fluoxetine can be prepared from the chiral alcohol intermediates (R)- or (S)-3-chloro-1-phenyl-1-propanol (CPPO), which could be produced via the enzymatic reduction of the carbonyl ketone substrate 3-chloro-1-phenyl-1-propanone (3-CPP; Scheme 1). Therefore, suitable reductase enzymes featuring chemo-, regio-, and stereoselectivity should be screened and employed for the efficient production of the target alcohol.
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Scheme 1

Reduction of 3-chloro-1-phenyl-1-propanone with reductase for the production of (R)- and (S)-3-chloro-1-phenyl-1-propanol, intermediates of fluoxetine

In this study, we generated ten different microbial reductases and screened these enzymes for their ability to produce a CPPO enantiomer. The properties of the carbonyl reductase thus selected, YOL151W, were characterized, and a reductase–glucose dehydrogenase coupling reaction was conducted in order to generate (S)-CPPO. Finally, a homology model of YOL151W and its docking model with cofactor and substrate were constructed to explain the detailed mechanism of the enantioselective reduction process.

Materials and methods

Chemicals

3-CPP, (R)- and (S)-CPPO, glucose dehydrogenase (GDH, Thermoplasma acidophilum), NAD(P)H, and NAD(P)+ were purchased from Sigma-Aldrich Co. (St. Louis, MO). All other chemicals were of analytical grade.

Cloning and expression of reductase gene

S. cerevisiae (KCTC 7904) chromosomal DNA was acquired from the Korea Research Institute of Bioscience and Biotechnology (KRIBB). The Leuconostoc citreum strain (KCTC 3721), which was supplied by the Biological Resource Center (Korea), was cultivated in lactobacilli formulations of deMan, Rogosa and Sharpe (MRS) broth at 30°C, and its genomic DNA was isolated by cell lysis and phenol/chloroform extraction. The Corynebacterium glutamicum (ATCC 13035) oxidoreductase gene (GenBank ID: NP-601323.1) was acquired from KRIBB.

Polymerase chain reaction (PCR) primer sets for yeast and Leuconostoc reductases were designed on the basis of nucleotide sequence information (Table 1) in the National Center for Biotechnology Information GenBank database. The PCR conditions were as follows: 30 cycles of 95°C for 1 min, 46°C for 1 min, and 72°C for 1 min. The PCR product was cloned into pGEM-T (Promega Corp., Madison, WI) and transformed into Escherichia coli XL1-Blue. The reductase gene was then ligated downstream of the T7 promoter of the pET-22b vector and used to transform E. coli BL21 (DE3). E. coli cells were cultured at 18°C in Luria–Bertani medium (1% tryptone, 0.5% yeast extract, and 0.5% NaCl) containing 100 μg/mL of ampicillin. When the cells reached an optical density600 nm of 0.5, expression was induced by the addition of 1 mM isopropyl thio-β-d-galactoside. Cultivation was continued for an additional 20 h, then the cells were harvested by centrifugation at 10,000×g for 10 min at 4°C and suspended in a 1/20 volume of 50 mM Tris–HCl buffer (pH 7.5). The cells were disrupted by sonication, and crude extracts were obtained by centrifugation at 10,000×g for 10 min.
Table 1

Screening of reductases suitable for the reduction of 3-chloro-1-pheny-1-propanone to 3-chloro-1-phenyl-1-propanol

Number

GenBank locus tag

Reported/putative function

Reductase activity (U/mL)

ΔNAD(P)Ha

ΔCPPOb

Saccharomyces cerevisiae

1

YJR096W

Aldo–keto reductase

1.18 (NADH)

ND

2

YDL124W

Alpha–keto amide reductase

0.85 (NADH)

ND

3

YOR120W

Glycerol dehydrogenase

0.21 (NADH)

ND

4

YGL185C

Hydroxy acid reductase

0.28 (NADH)

ND

5

YPL113C

Glyoxylate reductase

0.80 (NADH)

ND

6

YDR541C

Dehydrokaempferol 4-reductase

0.23 (NADPH)

ND

7

YCR107W

Aryl–alcohol reductase

0.35 (NADH)

ND

8

YOL151W

Methylglyoxal reductase

3.19 (NADPH)

0.55 (NADPH)

Leuconostoc citreum

9

LCK_01181

Oxidoreductase

0.30 (NADH)

ND

Corynebacterium glutamicum

10

NP-601323.1

Oxidoreductase

0.35 (NADPH)

ND

ND not detected

aReaction mixtures contained 1 mM 3-CPP, 0.2 mM NAD(P)H, and 50 μL cell-free extract in 1 mL of 50 mM Tris–HCl (pH 7.5). Spectrophotometric assay was used

bReaction mixtures contained 1 mM 3-CPP, 1 mM NAD(P)H, 0.2 U (spectrophotometrically assayed) cell-free extract in 2 mL of 50 mM Tris–HCl (pH 7.5). Reaction products were analyzed by HPLC. One unit of enzyme was defined as the amount of enzyme catalyzing the production of 1 μmol CPPO in 1 min at 30°C

Assay of reductase and glucose dehydrogenase activity

Reductase activity was assayed at 30°C by measuring the decrease in absorbance at 340 nm for 10 min using a spectrophotometer. The reaction mixture (1 mL) consisted of 1 mM 3-CPP (100 mM stock in DMSO), 0.2 mM NAD(P)H, 50 mM Tris–HCl buffer (pH 7.5), and 50 μL of cell-free extract. One unit of enzyme was defined as the quantity of enzyme required to catalyze the oxidation of 1 μmol NAD(P)H in 1 min at 30°C.

The glucose dehydrogenase oxidation reaction mixture (1 mL) consisted of 10 mM glucose, 2 mM NADP+, 50 mM Tris–HCl buffer (pH 7.5), and 0.05 μL of commercial glucose dehydrogenase. The reaction rate was monitored with a spectrophotometer on the basis of the increase in absorbance at 340 nm for 10 min at 30°C. One unit of enzyme was defined as the amount required to reduce 1 μmol of NADP+ in 1 min at 30°C.

Purification of reductase YOL151W

Reductase no. 8 (YOL151W) from baker's yeast was identified as an appropriate enzyme for the reduction of 3-CPP to CPPO in an enantioselective manner. The reductase protein in the cell extract was purified as follows. Firstly, the cell-free extract was loaded onto a Ni-NTA column (QIAGEN GmbH, Hilden, Germany) equilibrated with a 50-mM Tris–HCl buffer (pH 7.8) containing 50 mM imidazole and 300 mM NaCl. The recombinant reductase was then eluted from the column by the application of a 250-mM imidazole buffer. The active fractions were then collected and desalted with a PD-10 desalting column (GE Healthcare Bio-Sciences AB, UK).

Effects of temperature and pH

The effects of temperature and pH were evaluated using the purified YOL151W enzyme. Reactions were monitored spectrophotometrically by measuring the decrease in NADPH absorbance at 340 nm with 42.3 μg of enzyme. The reaction rate was measured at various temperatures (10–60°C). Meanwhile, in order to assess temperature stability, the enzyme was preincubated for 30 min at 10–50°C, then the remaining activity was assayed at 30°C. The following buffers (50 mM) were utilized to assess the effects of pH: pH 3–6, acetic acid/sodium acetate; pH 6–8, KH2PO4/K2HPO4; pH 7.5–10, Tris–HCl; pH 11–12, KCl–K2HPO4–K3PO4 buffers. In order to determine the pH stability, the enzyme was preincubated for 30 min in the pH buffers listed above at 0°C, then adjusted to pH 7.5, under which condition, the residual activity was determined.

Effect of buffer and DMSO concentrations

The effect of the concentration of Tris–HCl buffer (pH 7.5) on the reductase reaction rate was measured. The buffer was used at a concentration of 50, 100, 200, 250, or 300 mM and included 1 mM 3-CPP, 0.2 mM NADPH, and 10 μL of cell-free extract.

To evaluate the effect of DMSO, a reaction solution (100 mM Tris–HCl buffer, pH 7.5) containing 1 mM 3-CPP, 0.2 mM NADPH, 10 μL of cell-free extract, and DMSO at a final concentration of 1, 2, 3, 5, or 10% (v/v) was incubated, and the reaction rate was determined by observing the decrease in absorbance at 340 nm.

Enzymatic coupling reaction and reaction mixture analysis

The enantioselectivity of the enzymatic reduction of 3-CPP ketone was evaluated using an NADPH-recycling system. The general procedure was as follows: d-glucose (1.5-fold higher concentration than 3-CPP), 20 units of glucose dehydrogenase, 1 mM NADPH, various concentrations of 3-CPP in DMSO, and 10 units of reductase YOL151W were mixed in a total volume of 10 mL of 100 mM Tris–HCl buffer (pH 7.5), and the mixture was incubated at 30°C. The pH of the reaction mixture was monitored with a pH meter and maintained at 7.0–7.5 via the addition of 1 M NaOH. Four hundred microliter aliquots of the reaction mixture were sampled, mixed with 1.2 mL of ethyl acetate, and centrifuged at 10,000×g for 10 min. The upper phase containing ethyl acetate (1 mL) was obtained, combined with MgSO4, filtered, and then dried via vacuum pump centrifugation. The sample was resuspended in ethyl acetate (500 μL) and analyzed by a high-performance liquid chromatography (HPLC) system equipped with a CHIRALPAK IB column (Daicel Chemical Industries, Ltd., Tokyo, Japan). Separate peaks for the ketone substrate and (R)- and (S)-alcohols were obtained using n-hexane and 2-propanol (95:5, by volume) as the mobile phase at a flow rate of 0.8 mL/min. Detection was performed at 254 nm. Relative quantities were calculated on the basis of the peak area, which was suitably calibrated with standards of known concentration. Enantiomeric excess (ee) values were calculated from the alcohol products.

Computational methods

The initial 3D structural model of YOL151W was constructed based on the crystal structure of Sporobolomyces salmonicolor carbonyl reductase (SSCR; Protein Data Bank code: 1Y1P) (Kamitori et al. 2005). The target structure was modeled in silico by means of a structural homology search using HHpred/HHsearch (Söding et al. 2005) and homology modeling using MODELLER (Sali and Blundell 1993), then optimized using FoldX (Schymkowitz et al. 2005). The YOL151W active site was examined by comparison with the corresponding X-ray template of NADPH-bound SSCR. The geometry of the substrate (CPP) and cofactor (NADPH) was optimized using the Hartree–Fock method with a 6-31G* basis set as implemented in the Gaussian 03 program (Frisch et al. 2004). The restrained electrostatic potential procedure of the antechamber module from the AMBER suite was used to generate input files with charges for docking programs (Case et al. 2005). The result was used as a valid input for the AutoDock ligand preparation procedure.

Docking was performed with AutoDock (version 4.0), using the implemented empirical free energy function and the Lamarckian genetic algorithm (Huey et al. 2007). The best docked conformation was the one found to have the lowest binding energy and the greatest number of members in the cluster, indicating good convergence. The best orientation was identified and optimized using the scoring function based on the AMBER force field FF99 and energy minimization according to the Nelder and Mead algorithm (Case et al. 2005) for induced-fit simulation. The parameters embedded in the AMBER package were used for energy minimization and molecular dynamics, (Wang et al. 2004) and then, molecular dynamics (MD) simulations were performed. In addition, the docking of CPP and NADPH to our target protein was initially made according to predicted topological binding sites by several algorithms (Huang and Schroeder 2008).

Results

Screening for enantioselective reductases

Many microbial reductases have been isolated, and their genes are cloned, sequenced, and deposited in the GenBank database. In this study, we generated ten different microbial reductases in E. coli cells (Table 1). Eight of these originated from S. cerevisiae, and the remaining two enzymes were obtained from Leuconostoc and Corynebacterium species.

Although the expression levels of the recombinant enzymes were somewhat variable, the majority of target protein bands could be detected in soluble fractions on sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS–PAGE; see Supplementary data 1). The reductase activity in soluble fractions was determined via a spectrophotometric method according to the basic principle that NAD(P)H is oxidized when 3-CPP is reduced by the enzyme. Although the results differed according to which recombinant enzyme was tested, the majority evidenced measurable reductase activity (0.2–3.2 U/mL) toward 3-CPP (Table 1).

HPLC analysis was subsequently performed to detect the reaction product 3-CPPO. Among the enzymes tested, only reductase no. 8 produced a significant amount of (S)-CPPO.

Purification and characterization of reductase

Recombinant reductase no. 8 was purified in an effort to assess its reaction properties. As the enzyme harbored a His-tag at its C terminus, it could be readily purified via two sequential purification steps: Ni-NTA and PD-10 column chromatography. The purified enzyme had a specific activity of 0.99 U/mg toward the 3-CPP substrate.

The optimal reaction conditions for reductase no. 8 were evaluated via the spectrophotometric method. Analysis of the initial reaction rate demonstrated that the optimal temperature was 40°C, with activity decreasing rapidly at temperatures over 50°C (Fig. 1a). The enzyme was stable at up to 35°C for 30 min, while thermal stability decreased rapidly over 40°C (Fig. 1b). The optimal pH range was 7.5–8.0, and the enzyme was stable at pH 5–11 for 30 min (Fig. 1c, d).
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Fig. 1

Reaction properties of reductase no. 8. Reductase activity was determined by measuring the decrease in NADPH absorption at 340 nm. a Reductase activity measured at various temperatures. b Residual reductase activity measured after a 30-min preincubation at various temperatures. c Reductase activity measured at various pHs. d Residual reductase activity measured at pH 7.5 after a 30-min preincubation at various pHs. The buffers (50 mM) used in this experiments were acetic acid/sodium acetate (filled circles), KH2PO4–K2HPO4 (unfilled circles), Tris–HCl (filled squares), and KCl–K2HPO4–K3PO4 buffers (unfilled squares). e Effect of Tris buffer at concentrations of 50 (filled circles), 100 (unfilled circles), 200 (filled squares), 250 (triangles), and 300 mM (unfilled squares). f Effect of DMSO at concentrations of 1% (filled circles), 2% (unfilled squares), 3% (filled triangles), 5% (unfilled triangles), and 10% (squares)

In order to convert large quantities of substrate using a reductase, the reaction process should be coupled to a GDH enzyme reaction. The latter could regenerate NAD(P)H, a cofactor of the reductase. However, as this coupling reaction continues, glucose, a substrate of GDH, is oxidized into gluconic acid, and the solution pH is decreased, making the selection of buffer system critical. Therefore, the activity of reductase no. 8 was measured in an increasing molar concentration of Tris–HCl buffer. Reductase activity increased in concentrations of up to 100 mM, but decreased in concentrations over 200 mM (Fig. 1e).

In addition, since low substrate solubility presents another problem for the enzyme reaction, the substrate was dissolved in DMSO at a concentration of 1 M and then utilized for the reaction. Reductase activity was measured in increasing concentrations of DMSO in order to elucidate its effects (Fig. 1f). DMSO reduced enzyme activity only slightly up to a concentration of 3%, but its inhibitory effect increased significantly at concentrations over 5%.

The kinetic parameter of the reductase on 3-CPP and NADPH were measured (Table 2). Its kcat/Km values toward 3-CPP and NADPH were 163 and 260 min−1 mM−1, respectively.
Table 2

Kinetic parameters of YOL151W reductase

Ligand

kcat (min−1)

Km (mM)

kcat/Km (min−1 mM−1)

3-CPP

31.3

0.192

163

NADPH

29.1

0.112

260

Production of (S)-CPPO by reductase–GDH coupling reaction system

In order for the ketone reduction reaction to continue, sufficient NADPH must be supplied. Since this cofactor is a relatively expensive material, an NADPH regeneration system is required. Toward this end, we employed a GDH coupling reaction.

The products of the reductase–GDH coupling reaction were analyzed using an HPLC system equipped with a chiral column. As the reaction proceeded, the concentration of the ketone substrate decreased while the concentration of (S)-CPPO increased as a reaction product (Fig. 2). (R)-CPPO was not detected at all, implying that the e.e. value was 100%.
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Fig. 2

HPLC analysis of 3-CPP and CPPO enantiomer. a 3-CPP and (R)- and (S)-CPPO standard peaks. b Decrease in 3-CPP peak height and increase in (S)-CPPO peak height as a function of time. The reaction times were 15, 90, 120, and 180 min, respectively. The retention times of 3-CPP, (S)-CPPO, and (R)-CPPO were 7.8, 9.8, and 11.3 min, respectively

As the coupling reaction continued, gluconic acid accumulated, and the solution pH consequently decreased. The optimal pH of reductase no. 8 was 7.5–8.0, and its activity rapidly decreased at pH 6.0. In this experiment, the solution pH was maintained between pH 7.0–7.5 via occasional additions of a 1 M NaOH solution.

The coupling reaction was conducted with initial substrate concentrations of 10, 20, and 30 mM. Using 10 mM 3-CPP, all of the substrate was converted to (S)-CPPO within 60 min (Fig. 3a). When 20 and 30 mM 3-CPP were used, complete conversions were performed within approximately 300 and 360 min, respectively. When the substrate was supplied in three 10-mM aliquots, 30 mM (S)-CPPO was ultimately generated within 210 min (Fig. 3b). This result demonstrates that a higher conversion rate can be achieved by maintaining the substrate concentration below 10 mM.
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Fig. 3

Production of (S)-CPPO by reductase–GDH coupling reaction. a Time course of (S)-CPPO production using initial 3-CPP concentrations of 10 (filled circles), 20 (unfilled circles), and 30 mM (filled triangles). Enzyme reaction was performed in the absence of GDH (unfilled triangles). b Time course of (S)-CPPO production using three sequential additions of 3-CPP (10 mM each). Arrows indicate the times at which the second and third additions were made

Homology modeling and evaluation

By searching a similar structure for YOL151W using HHpred/HHsearch (Söding et al. 2005), we found an X-ray crystal structure, 1Y1P (Kamitori et al. 2005) as a potential homology template. The percent of sequence identity between the template and target was calculated to be 31.8% by Jotun–Hein method using DNAStar MegAlign program (see Supplementary data 2). We superimposed the active site of the target protein on the template (see Supplementary data 3).

The initial structure was refined by energy minimization and MD simulations. The total energy of the system was kept equilibrated after 150 ps. The average structures calculated during entire the 500 ps of MD simulations were adapted as the final model for YOL151W (Fig. 4a). The root mean square deviation (RMSD) value of overall main chain between YOL151W model structure and 1Y1P template structure was calculated to be 0.79 Å.
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Fig. 4

Homology model of YOL151W and its docking model. a Homology model constructed based on the crystal structure of SSCR (1Y1P). b CPP and NADPH docked to the enzyme active site. CPP is shown in gray, and NADPH is in yellow

The final model for YOL151W was assessed by Profile-3D (Luthy et al. 1992) and Procheck (Laskowski et al. 1993). Using Profile-3D, its compatibility score was 195 (the expected score for a protein of this size is 208). By employing Procheck, the reliability of the backbone torsion angles ψ–φ of the target protein was examined. The percentage of ψ–φ angles in the core Ramachandran region was 85.8%, which is comparable to that of template (89.7%). These data indicated that the 3D model (Fig. 4a) was reliable for further docking studies.

Docking of CPP to YOL151W

In order to explain the catalytic activity of YOL151W toward CPP in wet experiments, we performed a docking study (Fig. 4b). In the CPP–YOL151W complex, CPP is stabilized by hydrogen bonding and hydrophobic interactions in the center of the active site. Two hydrogen bonds are formed between them: the carbonyl oxygen of CPP binds to Ser127 and Tyr165. The complex has a favorable total interaction energy of −41.68 kcal/mol, in which the van der Waals and electrostatic energies are −33.09 and −8.59 kcal/mol, respectively. As seen in Fig. 4b, Phe132 and Met134 are positioned close to the benzyl ring of CPP, resulting in the second largest interaction energy, −3.35 kcal/mol (the strongest interactions come from the NADPH group), via π–π and sulfur–π interactions. Phe89 is also holding the ethyl chloride group of CPP via van der Waals interaction.

Discussion

(R)- and (S)-CPPO are chiral intermediates of antidepressant drugs including fluoxetine as mentioned above. These compounds could be generated from a prochiral ketone via an enantioselective reduction reaction, as depicted in Scheme 1. For the efficient production of (R)- and (S)-CPPO, a suitable reductase enzyme should be employed.

In this study, the reductase YOL151W was selected from among ten different microbial reductases for the production of (S)-CPPO. The gene GRE2 encoding YOL151W has previously been reported (Kayser et al. 2005) as have the substrate specificity and enantioselectivity of the reductase toward several compounds (Ema et al. 2006). Ema et al. reported YOL151W could produce ethyl-(R)-3-hydroxy-3-phenylpropanoate, an intermediate of fluoxetine with ee value of 70%. However, there has not yet been a report addressing the reactivity of the enzyme toward 3-CPP, the chiral intermediate of fluoxetine.

As our experimental data shows, reductase YOL151W exhibited both regiospecificity and enantioselectivity toward the carbonyl group of the 3-CPP molecule. The enzyme catalyzed the enantioselective reduction of the substrate, producing (S)-CPPO exclusively.

The reductase activity as measured by the spectrophotometric method was different from that obtained via HPLC (Table 1). We could not explain the exact reason why the different activities were observed, however, it seemed to be caused by the different assay systems; spectrophotometric assay (continuous assay) and HPLC assay (stop–sample, discontinuous assay).

In this study, homology modeling of YOL151W and docking modeling with CPP were performed to explain the mechanism of the enantioselective reduction of the substrate. As shown in Fig. 4, the carbonyl oxygen atom of CPP, located close to the nicotinamide ring, forms hydrogen bonds with Ser127 and Tyr165. The C4 atom of the nicotinamide ring possibly donates its hydrogen atom to the carbonyl carbon atom of CPP, with a distance of 3.1 Å between them. The proposed catalytic mechanism is as follows: The carbonyl oxygen atom of the substrate forms hydrogen bonds with both Tyr and Ser residues, and it is protonated by the Tyr residue, after which, a C4 hydrogen atom of NADPH attacks the carbonyl carbon atom of the substrate. Meanwhile, the nicotinamide mononucleotide ring of the cofactor is located on the re-face of CPP. Thus, this ketone should be reduced to (S)-CPPO (Fig. 5), which is consistent with our experimental observation.
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Fig. 5

Stereochemistry of the reductase YOL151W and a schematic representation of the reduction process

Some reductase reactions have occasionally been conducted using whole-cell systems. In this system, the reductase and glucose dehydrogenase enzymes were coexpressed in the same cell, thereby continuously recycling NAD(P)H (Ema et al. 2006; Moore et al. 2007). Additionally, various two-phase ionic liquid systems have been employed as reaction solvents to avoid substrate and/or product inhibition (Brautigam et al. 2007; Pfruender et al. 2006).

Collectively, the present research shows that, together with a glucose dehydrogenase coupling reaction, the recombinant yeast reductase YOL151W can be utilized to produce (S)-CPPO, and we might propose that it could be used to generate high concentrations of this product by adopting a whole-cell system or ionic liquid solvent system approach.

Acknowledgments

This work was supported by the 21C Frontier Microbial Genomics and Applications Center Program, Ministry of Education, Science and Technology, Republic of Korea and the 2009 Research Fund of the Catholic University of Korea.

Supplementary material

253_2010_2442_MOESM1_ESM.ppt (434 kb)
Supplementary data 1SDS–PAGE of expressed microbial reductases. Recombinant enzymes were expressed in E. coli BL21 (DE3) cells, and each of the soluble (S) and insoluble (P) fractions was analyzed. Arrows indicate the expected protein bands (PPT 434 kb)
253_2010_2442_MOESM2_ESM.ppt (246 kb)
Supplementary data 2Amino acid sequence and secondary structure of YOL151W aligned with those of SSCR. Blue arrows and orange bars indicate β-strands and α-helices, respectively (PPT 246 kb)
253_2010_2442_MOESM3_ESM.ppt (226 kb)
Supplementary data 33D model of YOL151W (green) superimposed on the X-ray crystal structure of SSCR (magenta) (PPT 226 kb)

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