Microbial Synthesis of Plant Oxylipins from γ-Linolenic Acid through

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Microbial Synthesis of Plant Oxylipins from γ‑Linolenic Acid through Designed Biotransformation Pathways Sae-Um Kim,† Kyoung-Rok Kim,§ Ji-Won Kim,† Soomin Kim,‡ Yong-Uk Kwon,‡ Deok-Kun Oh,§ and Jin-Byung Park*,† †

Department of Food Science and Engineering and ‡Department of Chemistry and Nano Science, Ewha Womans University, Seoul 120-750, Republic of Korea § Department of Bioscience and Biotechnology, Konkuk University, Seoul 143-701, Republic of Korea S Supporting Information *

ABSTRACT: Secondary metabolites of plants are often difficult to synthesize in high yields because of the large complexity of the biosynthetic pathways and challenges encountered in the functional expression of the required biosynthetic enzymes in microbial cells. In this study, the biosynthesis of plant oxylipinsa family of oxygenated unsaturated carboxylic acidswas explored to enable a high-yield production through a designed microbial synthetic system harboring a set of microbial enzymes (i.e., fatty acid double-bond hydratases, alcohol dehydrogenases, Baeyer−Villiger monooxygenases, and esterases) to produce a variety of unsaturated carboxylic acids from γ-linolenic acid. The whole cell system of the recombinant Escherichia coli efficiently produced (6Z,9Z)-12-hydroxydodeca-6,9-dienoic acid (7), (Z)-9-hydroxynon-6-enoic acid (15), (Z)-dec-4-enedioic acid (17), and (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid (2). This study demonstrated that various secondary metabolites of plants can be produced by implementing artificial biosynthetic pathways into whole-cell biocatalysis. KEYWORDS: plant oxylipins, γ-linolenic acid, biotransformation, whole-cell biocatalysis, Escherichia coli

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INTRODUCTION Biocatalysis is expanding its synthetic capacity by covering the preparation of large and complex molecules as well as small molecules with the help of the rapidly increasing number of enzymes discovered.1−6 Biocatalysis is especially useful in the synthesis of bioactive functional compounds (e.g., polyphenols, carotenoids, and terpenoids) found in nature, where their synthesis is hardly possible by any chemical means or reactions.5,7,8 Furthermore, there are products for which biological production remains challenging. These compounds include oxygenated unsaturated carboxylic acids that contain one or more hydroxyl or carboxyl groups as well as CC double bonds in the carbon skeleton such as oxylipins like as (6Z,9Z)12-hydroxydodeca-6,9-dienoic acid (7), (Z)-9-hydroxynon-6enoic acid (15), (4Z,7Z)-trideca-4,7-dienedioic acid (9), (Z)-dec-4-enedioic acid (17), or (6Z,9Z)-13-hydroxyoctadeca6,9-dienoic acid (2) (Figures 1 and 5). Oxygenated carboxylic acids are found in nature as components of diverse lipids (e.g., cerebrosides and waxes)9−13 and as secondary metabolites and signaling molecules (e.g., oxylipins).14,15 For instance, 11-hydroxyundec-9-enoic acid, 12-hydroxydodec-9-enoic acid, and 1,9-nonanedioic acid were identified as antifungal metabolites of the wild rice Oryza officinalis.14 12-Hydroxydodec-9-enoic acid was reported to stimulate the growth of the main root and lateral roots of plant seedlings.15 In view of industrial applications, ω-hydroxycarboxylic acids can be used for the production of α,ω-dicarboxylic acids and ω-aminocarboxylic acids, which are the starting materials of polyamides, polyesters, and other chemical products.16,17 In particular, a double bond in carboxylic acids allows for crosslinking or the addition of different functional groups to the © 2015 American Chemical Society

structure, which may lead to enhancements or innovations of different end products.18 Oxygenated carboxylic acids can be produced from the native plant biosynthetic pathways, which are composed of lipoxygenases, hydroperoxide lyases belonging to the CYP74 family, and alcohol dehydrogenases.15 However, the functional expression of the required biosynthetic enzymes in conventional microbial cells such as Escherichia coli or Saccharomyces cerevisiae was reported to be challenging.19,20 In the present study, we investigated oxygenation-based biocatalysis to produce a variety of oxygenated unsaturated carboxylic acids from a renewable biomass, which is difficult to obtain by traditional chemical means or reactions. On the basis of our previous proof-of-concept study, our approach was to design a biotransformation pathway consisting of fatty acid double-bond hydratases, alcohol dehydrogenases, Baeyer− Villiger monooxygenases (BVMOs), and esterases (Figures 1 and 5).21 Because most target products contain CC bond(s) in the carbon skeleton, γ-linolenic acid, which is abundant in nature, was used as the starting material.

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MATERIALS AND METHODS

Microbial Strains and Culture Media. The recombinant Escherichia coli ER2566 harboring the linoleate C12 double-bond hydratase gene from Lactobacillus acidophilus22 was grown at 37 °C with vigorous shaking in a Luria−Bertani (LB) medium supplemented with 50 μg mL−1 ampicillin. The expression of the recombinant Received: Revised: Accepted: Published: 2773

December 5, 2014 February 16, 2015 February 18, 2015 February 25, 2015 DOI: 10.1021/jf5058843 J. Agric. Food Chem. 2015, 63, 2773−2781

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Journal of Agricultural and Food Chemistry

Figure 1. Designed biotransformation pathway 1. γ-Linolenic acid (1) was converted into n-hexanoic acid (6) and (6Z,9Z)-12-hydroxydodeca-6,9dienoic acid (7) or n-pentanol (8) and (4Z,7Z)-trideca-4,7-dienedioic acid (9) by the multistep enzyme reactions. hydratase gene was induced by adding 0.1 mM isopropyl-β-Dthiogalactopyranoside (IPTG) at an optical density (OD600) of 0.5. Afterward, the culture was further incubated at 16 °C for 16 h. The recombinant E. coli BL21(DE3), used for hydration of γ-linolenic acid and biotransformation of hydroxy fatty acids, was cultivated in a Riesenberg medium, which was supplemented with 10 g L−1 glucose and the appropriate antibiotics for plasmid maintenance (Supporting Information Table S1). The Riesenberg medium consisted of 4 g L−1 (NH4)2HPO4, 13.5 g L−1 KH2PO4, 1.7 g L−1 citric acid, 1.4 g L−1 MgSO4, and 10 mL L−1 trace metal solution (10 g L−1 FeSO4, 2.25 g L−1 ZnSO4, 1.0 g L−1 CuSO4, 0.5 g L−1 MnSO4, 0.23 g L−1 Na2B4O7, 2.0 g L−1 CaCl2, and 0.1 g L−1 (NH4)6Mo7O24). Stenotrophomonas nitritireducens (Table S1) used for the hydration of the C9 double bond in γ-linolenic acid was cultivated in a nutrient medium as described previously.23 Reagents. γ-Linolenic acid (1), palmitic acid, n-hexanoic acid (6), n-pentanol (8), and ethyl acetate were obtained from Sigma-Aldrich. (6Z,9Z)-13-Hydroxyoctadeca-6,9-dienoic acid (2) and (6Z,12Z)-10hydroxyoctadeca-6,12-dienoic acid (10) were prepared via hydration of γ-linolenic acid with a linoleate C12 double-bond hydratase or a linoleate C9 double-bond hydratase in our laboratory (see Purification of Hydroxy Fatty Acids below for details). (6Z,9Z)-12-Hydroxydodeca-6,9-dienoic acid (7), (Z)-9-hydroxynon-6-enoic acid (15), (Z)dec-4-enedioic acid (17), and (4Z,7Z)-trideca-4,7-dienedioic acid (9) were synthesized from (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid (2) or (6Z,12Z)-10-hydroxyoctadeca-6,12-dienoic acid (10) via the recombinant E. coli-based biocatalysis in our laboratory (see Purification of Unsaturated Carboxylic Acids below for details). Purification of a Linoleate C12 Double-Bond Hydratase. The linoleate C12 double bond-hydratase of L. acidophilus was purified as previously reported.22 In brief, after cultivation of recombinant E. coli ER2566 harboring the linoleate C12 double-bond hydratase gene of L. acidophilus, the cell mass was harvested, washed, and resuspended into 50 mM phosphate buffer (pH 8.0) containing 300 mM NaCl, 10 mM imidazole, and 0.1 mM phenylmethanesulfonyl fluoride as a protease inhibitor. The resuspended cells were disrupted, and the

supernatant was applied to a His-Trap HP chromatography column (GE Healthcare) equilibrated with 50 mM phosphate buffer (pH 8.0) containing 300 mM NaCl. The active fractions were collected and dialyzed against 50 mM citrate−phosphate buffer (pH 7.0). Hydration of γ-Linolenic Acid. The enzymatic transformation of γ-linolenic acid into (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid was performed in 50 mM citrate−phosphate buffer (pH 7.0) containing 10 mM substrate and 2% (v/v) ethanol. In a 15 mL polypropylene tube with a screw cap, 3 mL of reaction mixture was incubated at 35 °C for 6 h with agitation at 200 rpm. The purified hydratases were applied to 3 mg mL−1. One unit (U) of the hydratase activity is defined as 1 μmol of (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid produced per minute at 35 °C. The whole-cell transformation of γ-linolenic acid into (6Z,9Z)-13hydroxyoctadeca-6,9-dienoic acid was performed in 50 mM Tris-HCl buffer (pH 7.0) containing 38 mM substrate and 0.05% (v/v) Tween 80. In a 15 mL polypropylene tube with a screw cap, 3 mL of reaction mixture was incubated at 35 °C for 24 h with agitation at 200 rpm. The recombinant E. coli BL21(DE3) expressing the linoleate C12 double-bond hydratase of L. acidophilus NBRC 13951 was added to 14.4 g L−1. The hydration of γ-linolenic acid into (6Z,12Z)-10-hydroxyoctadeca-6,12-dienoic acid with S. nitritireducens was carried out in 10 mL of 50 mM citrate−phosphate buffer (pH 7.0). γ-Linolenic acid was added to a reaction medium containing 20 g dry cells L−1 and 0.05% (v/v) Tween 80. The reaction mixture was incubated at 30 °C and sparged with a gas mixture consisting of 95% N2 and 5% CO2 at 0.02 vvm to maintain anaerobic conditions.23 Biotransformation of Hydroxy Fatty Acids. The biotransformation of the hydroxy fatty acids with recombinant E. coli cells was carried out on the basis of our earlier work.21 Briefly, the recombinant cells were cultivated in the Riesenberg medium at 30 °C, and the expression of the target genes was induced with 0.1 mM IPTG and/or 2.0 g L−1 rhamnose at an OD600 of 0.5. Thereafter, the cultivation temperature was reduced to 20 °C to facilitate the soluble expression of the target genes (Supporting Information Figure S4). When the added 2774

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Journal of Agricultural and Food Chemistry glucose was consumed, the recombinant cells were harvested by centrifugation at 3500g for 15 min at 4 °C and used as biocatalysts. The biotransformation was initiated by adding 2 mM of a hydroxy fatty acid to 50 mM Tris-HCl buffer (pH 8.0) containing 0.5 g L−1 Tween 80 and 7.2 g dry cells L−1 of the recombinant E. coli BL21(DE3) that expressed the ADH from Micrococcus luteus, the esterase from P. f luorescens SIK WI, and the BVMO from either Pseudomonas putida KT2440 or Pseudomonas f luorescens DSM 50106. The biotransformations were conducted in a shaking incubator (35 °C and 200 rpm). Product Analysis by GC-MS. The concentrations of the remaining fatty acids and accumulating carboxylic acids in the medium (i.e., γ-linolenic acid (1), (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid (2), (6Z,9Z)-12-hydroxydodeca-6,9-dienoic acid (7), (Z)-9-hydroxynon-6-enoic acid (15), (4Z,7Z)-trideca-4,7-dienedioic acid (9), and (Z)-dec-4-enedioic acid (17)) were determined as described previously.21 The reaction medium was mixed with an equal volume of ethyl acetate containing 2 or 5 g L−1 palmitic acid as an internal standard. The organic phase was harvested after vigorous vortexing and then derivatized with N-methyl-N-(trimethylsilyl)trifluoroacetamide (MSTFA). The trimethylsilyl (TMS) derivatives were analyzed using a Thermo Ultra Trace GC system connected to an ion trap mass detector (Thermo ITQ1100 GC-ion Trap MS; Thermo Scientific). The derivatives were separated on a nonpolar capillary column (30 m length, 0.25 μm film thickness, HP-5MS; Agilent, Santa Clara, CA, USA). A linear temperature gradient was programmed as follows: 90 °C, 15 °C min−1 to 200 °C; 200 °C, 5 °C min−1 to 250 °C; 90 °C, 15 °C min−1 to 200 °C; and 200 °C, 5 °C min−1 to 280 °C. The injection port temperature was 230 °C. Mass spectra were obtained by electron impact ionization at 70 eV. Scan spectra were obtained within the range of m/z 100−600. Selected ion monitoring (SIM) was used for the detection and fragmentation analysis of the reaction products. Product Analysis by GC. The obtained hydroxy fatty acids were derivatized by using a 3:1 mixture of pyridine and MSTFA. TMSderivatized hydroxy fatty acids were analyzed by GC (Agilent 6890N) equipped with a flame ionization detector and an SPB-1 capillary column (15 m × 0.32 mm inside diameter, 0.25 μm thickness; Supelco) using palmitic acid as an internal standard.23 The column temperature increased from 150 to 210 °C at an initial rate of 4 °C min−1 and then at rate of 30 °C min−1 until 280 °C, and this temperature was maintained at 280 °C for 5 min. The injector and detector temperatures were held at 260 and 250 °C, respectively. Identification of the Biotransformation Products. The biotransformation products were identified by GC-MS or NMR analyses. (Z)-9-Hydroxynon-6-enoic acid (15), (4Z,7Z)-trideca-4,7dienedioic acid (9), and (Z)-dec-4-enedioic acid (17) were identified by GC-MS analysis and comparison to pure reference compounds (Figures 4 and 7). (6Z,9Z)-13-Hydroxyoctadeca-6,9-dienoic acid (2) and (6Z,9Z)-12-hydroxydodeca-6,9-dienoic acid (7) were identified by GC-MS and NMR analyses (see Figure S3 in the Supporting Information for identification of (6Z,9Z)-13-hydroxyoctadeca-6,9dienoic acid and Figure S5 for identification of (6Z,9Z)-12hydroxydodeca-6,9-dienoic acid). Identification of compounds 2, 7, 9, 15, and 17 was further confirmed by the GC-MS analysis of their hydrogenated products, which were obtained by catalytic hydrogenation (Figures 4, 7, and S1B). Purification of the Biotransformation Products. Purification of Hydroxy Fatty Acids. The hydroxy fatty acids (i.e., (6Z,9Z)-13hydroxyoctadeca-6,9-dienoic acid (2) and (6Z,12Z)-10-hydroxyoctadeca-6,12-dienoic acid (10)) were isolated from the reaction medium by solvent extraction and column chromatography as described previously.23 In brief, the biotransformation products were isolated by the extraction using two volumes of ethyl acetate two times. After the organic layer was separated from the aqueous phase, the organic extracts were dried with Na2SO4 and concentrated by the evaporation in vacuo. The concentrate was purified by the silicic acid column chromatography with methanol gradient (0−5%, v/v) in CH3Cl (50 mL). Any impurities and the remaining substrate were washed off with CH3Cl and 1% methanol in CH3Cl. In the case of (6Z,9Z)-13hydroxyoctadeca-6,9-dienoic acid, the concentrate was purified with a semipreparative HPLC system (Agilent) equipped with a Nucleosil

Table 1. Products Accessible through the Multistep Biocatalysis starting material

final product

product yielda (%)

γ-linolenic acid (1)

(6Z,9Z)-13-hydroxyoctadeca6,9-dienoic acid (2) (6Z,12Z)-10hydroxyoctadeca-6,12dienoic acid (10)

92 ± 2 (60%, 96%)b 97 ± 4

(6Z,9Z)-13-hydroxyoctadeca6,9-dienoic acid (2)

(6Z,9Z)-12-hydroxydodeca6,9-dienoic acid (7) (4Z,7Z)-trideca-4,7dienedioic acid (9)

81 ± 4 (43%, 71%)b 34 ± 6c

(6Z,12Z)-10hydroxyoctadeca-6,12dienoic acid (10)

(Z)-9-hydroxynon-6-enoic acid (15) (Z)-dec-4-enedioic acid (17)

72 ± 7 73 ± 4

a

The product yield was calculated on the basis of the substrate depletion and the product concentration, which were determined by GC-MS. All experiments were carried out in triplicate. bThe numbers in parentheses indicate the isolated yield and purity, respectively. The isolated yields were calculated on the basis of the amount of the products isolated and the amount of the substrates added into the reaction medium. cThe product yields were low compared to those of the other products because (6Z,9Z)-12-hydroxydodeca-6,9-dienoic acid (7) was coproduced during biotransformation (Figure 3B). C-18 semiprep column as described in a previous study24 (see Figure S2 in the Supporting Information for purification of (6Z,9Z)13-hydroxyoctadeca-6,9-dienoic acid). Purification of Unsaturated Carboxylic Acids. The medium-chain, unsaturated carboxylic acids (e.g., 12-hydroxydodeca-6,9-dienoic acid, 9-hydroxynona-6-enoic acid, and 1,10-deca-6-enedioic acid) were isolated from the whole-cell reaction medium as described previously.21,25 In brief, the biotransformation products were extracted with ethyl acetate three times. The extract was dried over MgSO4 and concentrated via evaporation in vacuo. The impurities (e.g., n-alkanoic acids coproduced during biotransformations and long-chain fatty acids originating from the starting materials) were removed by washing with hexane (see Figure S6 in the Supporting Information for purification of (6Z,9Z)-12-hydroxydodeca-6,9-dienoic acid). (6Z,9Z)-12-Hydroxydodeca-6,9-dienoic acid was also purified by HPLC (Figure S5).

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RESULTS The fatty acid double-bond hydratases and BVMOs were the key enzymes in our designed biotransformation pathways because the structures of the final products were determined by the regioselectivity of both enzymes (Figures 1 and 5). Thereby, the first step was to select the appropriate enzymes in terms of substrate scope and regioselectivity for the production of the target compounds. Screening of a γ-Linolenic Acid C12 Double-Bond Hydratase. Recently, a linoleate C12 double-bond hydratase of L. acidophilus NBRC 13951 was cloned and characterized.22 Thereby, the enzyme was first examined for hydration of the C12 double bond in γ-linolenic acid. The enzyme was produced in E. coli ER2566 and subjected to purification via His-Trap affinity chromatography. Afterward, the purified enzyme was added to 50 mM citrate−phosphate buffer (pH 7.0) containing 10 mM γ-linolenic acid. A single product was accumulated in the reaction mixture (Figure 2A and Supporting Information Figure S1A). The product (2) was purified to a purity of >96% via solvent extraction and column chromatography (Figure S2). The mass spectrum contained peaks at m/z 173 and 369 due to the α-cleavages at both sides of the hydroxy-TMS function at 2775

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Figure 3. Biotransformation of (6Z,9Z)-13-hydroxyoctadeca-6,9dienoic acid. (6Z,9Z)-13-Hydroxyoctadeca-6,9-dienoic acid (2) was converted into (A) (6Z,9Z)-12-hydroxydodeca-6,9-dienoic acid (7) or (B) (6Z,9Z)-12-hydroxydodeca-6,9-dienoic acid and (4Z,7Z)-trideca4,7-dienedioic acid (9). The first biotransformation was conducted with the recombinant E. coli BL21(DE3) pACYC-ADH, pET-BmoF1, pCOLA-PFE1 expressing the ADH from M. luteus, the BVMO from P. fluorescens DSM 50106, and the esterase from P. f luorescens SIK WI. The second biotransformation was conducted with the recombinant E. coli BL21(DE3) pACYC-ADH-BVMO, pCOLA-PFE1, expressing the ADH from M. luteus, the BVMO from Ps. putida KT2440, and the esterase from P. f luorescens SIK WI. The experiments were carried out in triplicate, and the error bars indicate the standard deviation.

Figure 2. Biotransformation of γ-linolenic acid. γ-Linolenic acid (1) was converted into (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid (2) by (A) the isolated linoleate C12 double-bond hydratase of L. acidiphilus NBRC 13951 or (B) the recombinant E. coli BL21(DE3) expressing the linoleate C12 double-bond hydratase of L. acidiphilus NBRC 13951. The experiments were carried out in triplicate, and the error bars indicate the standard deviation.

C13, whereas the mass spectrum of the hydrogenation product showed peaks at m/z 173 and 373 (Figure S1B). The resonance signals at 3.63 ppm (m, 1H) and four olefinic protons at 5.38 (m, 4H, H-6, H-7, H-9, and H-10) in 1H NMR indicated the existence of a hydroxyl group and a double bond (Figure S3). The observed signal at 71.8 ppm (C-13) of 13C NMR also supported the existence of one hydroxyl group (Figure S3). On the basis of these data, the product was identified as (6Z,9Z)13-hydroxyoctadeca-6,9-dienoic acid (2). The concentration of (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid in the reaction medium increased to 6 mM. The hydration rate was increased to 21 U g protein−1 at t < 1 h. The isolated yield of the compound was ∼60% with a purity of 96% (Table 1). This result indicated that the linoleate C12 double-bond hydratase also exhibited very high regioselectivity with γ-linolenic acid. Whole-Cell Production of (6Z,9Z)-13-Hydroxyoctadeca-6,9-dienoic Acid. The microbial production of (6Z,9Z)13-hydroxyoctadeca-6,9-dienoic acid (2) from γ-linolenic acid was also investigated. After batch cultivation of recombinant E. coli BL21(DE3) expressing the linoleate C12 double-bond

hydratase of L. acidophilus NBRC 13951, cell harvest via centrifugation, and resuspension into 50 mM Tris-HCl buffer (pH 7.0), γ-linolenic acid was added to a concentration of 38 mM in the reaction medium. γ-Linolenic acid was converted into the target product at a rate of 2.9 μmol g dry cells−1 min−1 (i.e., 2.9 U g dry cells−1) at t < 10 h, thus resulting in a final product concentration of ∼30 mM (8.9 g L−1) in the reaction medium (Figure 2B). The bioconversion yield, which was calculated on the basis of the substrate depletion and the product concentration determined by GC-MS, reached >95%. The byproduct formation was hardly observed (Supporting Information Figure S1B). These results indicated that the whole-cell biotransformation system was quite efficient in terms of reaction selectivity and rate. Biotransformation of (6Z,9Z)-13-Hydroxyoctadeca6,9-dienoic Acid (2). Hydroxy fatty acids are used as starting materials for the production of diverse secondary metabolites and signaling molecules in living organisms.12,13 Thus, we 2776

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Figure 4. GC-MS analysis of (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid biotransformation products. (A) GC-MS of the biotransformation products, which were produced with the recombinant E. coli BL21(DE3) pACYC-ADH, pET-BmoF1, pCOLA-PFE1. The bottom panel shows the mass spectrum of the hydrogenated product of (6Z,9Z)-12-hydroxydodeca-6,9-dienoic acid, which is identical to that of the authentic compound 12-hydroxydodecanoic acid. (B) GC-MS of the biotransformation products, which were produced with the recombinant E. coli BL21(DE3) pACYCADH-BVMO, pCOLA-PFE1. The bottom panel shows the mass spectrum of the hydrogenated product of (4Z,7Z)-trideca-4,7-dienedioic acid, which is identical to that of the authentic compound tridecane-1,13-dioic acid.

Figure 5. Designed biotransformation pathway 2. γ-Linolenic acid (1) is converted into 2-nonenoic acid (14) and (Z)-9-hydroxynon-6-enoic acid (15) or 2-octenol (16) and (Z)-dec-4-enedioic acid (17) by the multistep enzyme reactions.

investigated the biotransformation of (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid (2) into C12 oxylipins (e.g., (6Z,9Z)-12-

hydroxydodeca-6,9-dienoic acid (7)). The substrate (2) was prepared after the whole-cell reaction of γ-linolenic acid, 2777

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Journal of Agricultural and Food Chemistry isolation via ethyl acetate extraction, and concentration via evaporation in vacuo. The BVMO enzymes, which determine the chemical structure of the final products due to their regioselectivity, were first examined. The BVMOs that are active with long-chain aliphatic ketones26−29 were chosen as candidates. When the recombinant E. coli BL21(DE3) pACYCADH, pET-BmoF1, pCOLA-PFE1 expressing an alcohol dehydrogenase (ADH) from Micrococcus luteus NCTC2665, a BVMO from Pseudomonas fluorescens DSM 50106,30 and an esterase from P. f luorescens SIK WI was added to the reaction medium containing (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid (2), the compound was selectively converted into (6Z,9Z)-12-hydroxydodeca-6,9-dienoic acid (7) and n-hexanoic acid (6) via (6Z,9Z)-13-oxooctadeca-6,9-dienoic acid (3) and the ester (4) (Figures 3A and 4A). (6Z,9Z)-12-Hydroxydodeca-6,9-dienoic acid (7) was identified via GC-MS and NMR analyses after purification (Figures 4A and S5). Notably, the target product (7) was produced with a final concentration of 1.6 mM from 2.0 mM of the substrate (6Z,9Z)-13hydroxyoctadeca-6,9-dienoic acid. On the other hand, the addition of the recombinant E. coli BL21(DE3) pACYC-ADH-BVMO, pCOLA-PFE1 expressing an ADH from M. luteus NCTC2665, a BVMO from Pseudomonas putida KT2440,27 and an esterase from P. f luorescens SIK WI resulted in a production of not only (6Z,9Z)-12-hydroxydodeca6,9-dienoic acid (7) and n-hexanoic acid (6) but also (4Z,7Z)trideca-4,7-dienedioic acid (9) and n-pentanol (8) (Figures 3B and 4B). (4Z,7Z)-Trideca-4,7-dienedioic acid was identified on the basis of the GC-MS analysis data of the pure compound 1,12dodec-9-enedioic acid,31 which has the same functional group in the fatty acid skeleton. It was assumed that the ester compound (5) was produced via the formation of the abnormal ester, which was driven by migration of the less-substituted carbon center during BVMO catalysis.32,33 Biotransformation of (6Z,12Z)-10-Hydroxyoctadeca6,12-dienoic Acid (10). The biotransformation of (6Z,12Z)10-hydroxyoctadeca-6,12-dienoic acid (10) (Figure 5), which had been produced from γ-linolenic acid by using the hydratase of S. nitritireducens23 (Supporting Information Figure S7), was also carried out using the setup identical to the biotransformation of (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid (2). (Z)9-Hydroxynon-6-enoic acid (15) and 2-nonenoic acid (14) were produced as the major products by recombinant E. coli BL21(DE3) pACYC-ADH-BVMO, pCOLA-PFE1 (Figures 6A and 7A). 9-Hydroxynona-6-enoic acid was identified on the basis of the GC-MS analysis data of the pure compound 12hydroxyundec-9-enoic acid,15 which has the same functional group in the fatty acid skeleton. Notably, the BVMO of P. putida KT2440 showed quite high selectivity with oxygenation of (6Z,12Z)-10-oxooctadeca-6,12-dienoic acid (11), but it showed low selectivity with (6Z,9Z)-13-oxooctadeca-6,9dienoic acid (3) (Figure 3B). The biotransformation of (6Z,12Z)-10-hydroxyoctadeca6,12-dienoic acid (10) with recombinant E. coli BL21(DE3) pACYC-ADH, pET-BmoF1, pCOLA-PFE1 expressing the BVMO from P. f luorescens DSM50106 instead of the BVMO from P. putida KT2440 resulted in the production of (Z)-dec-4-enedioic acid (17) and 2-octenol as the major products (Figures 6B and 7B). These results indicated that a variety of plant oxylipins could be produced via combined biocatalysis using fatty acid double-bond hydratases and BVMOs.

Figure 6. Biotransformation of (6Z,12Z)-10-hydroxyoctadeca-6,12dienoic acid. (6Z,12Z)-10-Hydroxyoctadeca-6,12-dienoic acid (10) was converted by (A) the recombinant E. coli BL21(DE3) pACYCADH-BVMO, pCOLA-PFE1, and (B) the recombinant E. coli BL21(DE3) pACYC-ADH, pET-BmoF1, pCOLA-PFE1. The experiments were carried out in triplicate, and the error bars indicate standard deviation.

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DISCUSSION A family of oxygenated unsaturated carboxylic acids (e.g., oxylipins) that play important roles in plant signaling and defense are usually produced via a serial reaction of lipoxygenases and hydroperoxide lyases belonging to CYP74 family.15 For instance, 12-hydroxydodec9-enoic acid, 12-oxododec-9-enoic acid, and 1,12-dodec-9-enedioic acid, which were reported to stimulate growth of the roots of plant seedlings, were synthesized from α-linolenic acid by 13lipoxygenases, hydroperoxide lyases, and alcohol dehydrogenases. 9-Hydroxynonanoic acid and 1,9-nonanedioic acid, which are involved in defense against infection, were produced from α-linolenic acid by 9-lipoxygenases, hydroperoxide lyases, and alcohol dehydrogenases.15 Because the oxygenated carboxylic acids have a great potential not only as agrochemicals but also as building blocks and/or additives for the production of plastics (e.g., polyamides and polyesters), resins, hot melt adhesives, powder coatings, corrosion inhibitors, lubricants, plasticizers, greases, and perfumes,14,18,34,35 their biosyntheses have been extensively investigated. For instance, 9-oxononanoic acid was produced from linoleic acid via an enzymatic two-step reaction catalyzed by the 9S-lipoxygenase of Solanum tuberosum and the 9/13-hydroperoxide lyase of Cucumis melo.19 Another example was the biotransformation of α-linoleic acid into 3(Z)-hexenal 2778

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Figure 7. GC-MS analysis of (6Z,12Z)-10-hydroxyoctadeca-6,12-dienoic acid biotransformation products. (A) GC-MS of the biotransformation products, which were produced with the recombinant E. coli BL21(DE3) pACYC-ADH-BVMO, pCOLA-PFE1. The bottom panel shows the mass spectrum of the hydrogenated product of (Z)-9-hydroxynon-6-enoic acid, which is identical to that of the authentic compound 9-hydroxynonanoic acid. (B) GC-MS of the biotransformation products, which were produced with the recombinant E. coli BL21(DE3) pACYC-ADH, pET-BmoF1, pCOLA-PFE1. The bottom panel shows the mass spectrum of the hydrogenated product of (Z)-dec-4-enedioic acid, which is identical to that of the authentic compound decane-1,10-dioic acid.

small number of BVMOs including the BVMO from P. f luorescens DSM 50106,30 P. putida KT2440,27,39 R. jostii RHA1,40 P. veronii MEK700,26 P. cepacia,28 and Mycobacterium tuberculosis29 were found to be active with aliphatic ketones. The present study herein demonstrated that product profiles of the novel biosynthetic pathways (Figures 1 and 5) were dependent on the catalytic activity, but they were especially dependent on the regioselectivity of the BVMOs (Figures 3 and 6). The BVMO originating from P. f luorescens DSM 50106 catalyzed the normal ester formation with (6Z,9Z)-13-oxooctadeca-6,9dienoic acid (3), which was driven by migration of the moresubstituted carbon center during BVMO catalysis (Figure 3A). However, the BVMO from P. putida KT2440 showed a quite high activity toward the abnormal ester formation32,33 in the case of (6Z,9Z)-13-oxooctadeca-6,9-dienoic acid (3) (Figure 3B). Notably, the BVMO from P. f luorescens DSM 50106 catalyzed the abnormal ester formation of (6Z,12Z)-10-oxooctadeca-6,12dienoic acid (11), thus resulting in the production of 1,10dec-6-enedioic acid (Figure 6B). Unfortunately, the BVMOs from P. f luorescens DSM 50106 and P. putida KT2440 were too unstable to allow for the proper investigation of their kinetic properties and their 3-D structures. As a result, a detailed investigation of their regioselectivity is currently severely hampered. In summary, we demonstrated that a variety of plant oxylipins (i.e., (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid (2), (6Z,9Z)-12-hydroxydodeca-6,9-dienoic acid (7), (Z)-9-hydroxynon-6-enoic acid (15), (4Z,7Z)-trideca-4,7-dienedioic acid (9), and (Z)-dec-4-enedioic acid (17)) could be produced from a renewable fatty acid (i.e., γ-linolenic acid) via the regioselective BVMO-based whole-cell biocatalysis. This study will contribute to the preparation of industrially relevant unsaturated carboxylic acids.

and 12-oxododec-9-enoic acid by a recombinant S. cerevisiae expressing a soybean lipoxygenase and watermelon hydroperoxide lyase.20 However, the productivity of the biocatalysts appeared to be limited due to the low expression level of the plant enzymes in the microbial cells. This study herein demonstrated that a variety of the industrially relevant fatty acids and carboxylic acids (i.e., (6Z,9Z)-13-hydroxyoctadeca-6,9-dienoic acid (2), (6Z,9Z)-12hydroxydodeca-6,9-dienoic acid (7), (Z)-9-hydroxynon-6-enoic acid (15), (4Z,7Z)-trideca-4,7-dienedioic acid (9), and (Z)-dec4-enedioic acid (17)) could be produced from renewable fatty acids (i.e., γ-linolenic acid) by using artificial multistep enzyme reaction pathways (Figures 1 and 5). The synthetic pathways were composed of enzymes originating solely from bacteria, which could be overexpressed easily in functional forms in bacterial cells. Some of the key enzymes in the biotransformations reported herein are the BVMOs. The regiospecificity of the enzymes was crucial for providing access to the desired reaction products (Figures 1 and 5). The BVMOs (EC 1.4.13.22) catalyzed the insertion of oxygen into the carbon skeletons adjacent to the carbonyl groups.36−38 The regiochemistry of the reaction was governed by predictable conformational, steric, and electronic effects as the rearrangement process of the tetrahedral peroxo Criegee intermediate proceeded with strict retention of configuration. Common representative substrates were simple cyclic ketones (e.g., cyclobutanone, cyclopentanone, cyclohexanone, and cycloheptanone), chiral cyclic ketones (e.g., 2-methylcyclopentanone and bicyclo[3.2.0]hept-2-en-6-one), and aliphatic ketones (e.g., 2-octanone).38 Most BVMOs discovered so far showed catalytic activity for the oxygenation of simple cyclic ketones and chiral cyclic ketones, and only a 2779

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ASSOCIATED CONTENT

S Supporting Information *

Table S1 and Figures S1−S7. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*(J.-B.P.) E-mail: [email protected]. Funding

This study was supported by a Marine Biomaterials Research Center grant from the Marine Biotechnology Program funded by the Ministry of Oceans and Fisheries, Korea. Notes

The authors declare no competing financial interest.

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