Cloning and characterization of enoate reductase with high β-ionone to dihydro-β-ionone bioconversion productivity
- Xuesong Zhang†1, 3,
- Shiyong Liao†2,
- Fuliang Cao1,
- Linguo Zhao1, 2Email author,
- Jianjun Pei2, 4 and
- Feng Tang5
© The Author(s). 2018
Received: 11 October 2017
Accepted: 23 April 2018
Published: 9 May 2018
Dihydro-β-ionone is a principal aroma compound and has received considerable attention by flavor and fragrance industry. The traditional method of preparing dihydro-β-ionone has many drawbacks, which has restricted its industrial application. Therefore, it is necessary to find a biotechnological method to produce dihydro-β-ionone.
In this study, the enoate reductase with high conversion efficiency of β-ionone to dihydro-β-ionone, DBR1, was obtained by screening four genetically engineered bacteria. The product, dihydro-β-ionone, was analyzed by GC and GC-MS. The highest dihydro-β-ionone production with 308.3 mg/L was detected in the recombinant strain expressing DBR1 which was later on expressed and purified. Its optimal temperature and pH were 45 °C and 6.5, respectively. The greatest activity of the purified enzyme was 356.39 U/mg using β-ionone as substrate. In the enzymatic conversion system, 1 mM of β-ionone was transformed into 91.08 mg/L of dihydro-β-ionone with 93.80% of molar conversion.
DBR1 had high selectivity to hydrogenated the 10,11-unsaturated double bond of β-ionone as well as high catalytic efficiency for the conversion of β-ionone to dihydro-β-ionone. It is the first report on the bioconversion of β-ionone to dihydro-β-ionone by using enoate reductase.
Ionones were widely used in daily flavor formula. Most Ionons were ionon and methyl ionone, and it cannot satisfy the market demand with weak production capacity and single variety. Dihydro-β-ionone, also called ‘sweet osmanthus king’, has received great attention from the flavour and fragrance industry. It is a main aroma compound with mellow, sweet, and fresh cedar scent in Osmanthus oil. Because of its unique scent, commercial extracts are in high demand for use in the production of expensive perfumes and cosmetics . It is widely used in foodstuff and beverage industries. It is also an important intermediate compound used in the synthesis of tea screw alkanes and its analogues with great application foreground [2, 3]. Dihydro-β-ionone naturally occurs in Osmanthus fragrans Lour, roses, and in many flowers. For a number of ingredients, their extractions from plant or animal tissues are the best way to get the products. However, the aroma compound dihydro-β-ionone is of particular interest because it occurs mostly at low concentrations in natural tissues and extraction in most situations is not economically feasible . In addition, the extraction of aroma compounds from natural source is an expensive and arduous task and strongly depends on agriculture and all the factors surrounding it . The chemical production process of dihydro-β-ionone is a catalyzed hydrogenation process from β-ionone, which is performed by using asymmetric hydrogenation of costly catalysts like chiral rhodium or ruthenium phosphines [6, 7]. The limiting issues involved in the chemical production process are the disposal of complex heavy metal ligands and the requirement of high pressure . Therefore, a specific branch of biotechnology has been developed in which constraints relative to the natural state are present .
Biotechnology represents a very attractive alternative for the sustainable production of flavors and fragrances . With the increasing development of genetic engineering, it became possible to produce heterologous products in microbial cell factories that are normally found only in small amounts in nature [11, 12]. The highly demanded phenethyl alcohol, vanillin, and γ-decalactone can also be produced through biotechnology of natural products, but, for many compounds, yields are very low and the products are too expensive to satisfy demand [13–17]. In recent years, enoate reductases which are members of the ‘old yellow enzyme family’, have been attracting the interest of chemists for its high potential in asymmetric reduction of activated C=C bonds . Enoate reductases have a wide variety of substrates, such as α,β-unsaturated aldehyde ketone, nitroalkenes, α,β-unsaturated nitriles, α,β-unsaturated carboxylic acids and their derivatives [19–21]. Enoate reductases have been found in plant, bacteria, fungi, and protozoa [22–25], and play critical roles not only in the biosynthesis of steroids, fatty acids, and phytohormones like jasmonic acid , but also in plant secondary metabolism, such as pulegone and artemisinin biosynthesis [27, 28]. In the area of enoate reductases enzymatic hydrogenation, research efforts mainly focused on whole-cell systems with wild-type microorganisms, new enoate reductases with highly catalytic activity, and broad substrate profile . There exists almost no research on the conversion of natural active substance with enoate reductases, and the demand of industrial production could not be met because of the absence of high activity, stability, and tolerance to the system.
In the present study, high enoate reductases were screened and β-ionone was converted into dihydro-β-ionone by whole-cell systems. Overpression and characterization of the optimization of enoate reductases have also been reported. This enzyme has high selectivity for transforming β-ionone to dihydro-β-ionone. These extraordinary properties enable enoate reductases a good biocatalyst for producing dihydro-β-ionone in vitro.
Bacterial strains and plasmids
The genes encoding DBR2 (NCBI accession number BAU61367.1) from Artemisia annua, BacOYE1 (NCBI accession number KJ577134.1) from Balillus sp., DBR1 (NCBI accession number FJ750460.1) from Artemisia annua, and Unigene (CL2687.Contig2_ALL) from Osmanthus fragrans transcriptome sequencing (SRA accession number SRP057917) which shared 78% identity with the DBR2 of Artemisia annua (NCBI accession number BAU61367.1) were synthesized by Shanghai Generay Biotech Co. Ltd. (Shanghai, China). The synthetic genes were inserted into plasmid PGEX-4 T1 or pET-28a (Invitrogen, USA) to generate the expression vector pET-28a-BacOYE1, pET-28a-DBR1, PGEX-4 T1-DBR2, and PGEX-4 T1-2687. Then recombinant plasmids were transformed into E. coli BL21 (DE3). The sequence of DBR1, DBR2, BacOYE1, and CL2687.Contig2_ALL was shown in Additional file 1.
Production of dihydro-β-ionone on whole-cell systems
One hundred microliters of an overnight culture was used to inoculate 5 ml of LB resistance medium at 37 °C to an optical density at OD600 of 0.4–2.0. The expression of the proteins was induced by adding 0–1 mM isopropyl-b-D-thiogalactoside (IPTG). Fifty to three hundred microliters β-ionone solution to a final concentration of 10 mM was added using chloroform, DMSO, methanol, ethanol and ethyl acetate as the solvents. The bacteria were further incubated at 22–42 °C for 6–27 h and gently shaken (180 rpm). One milliliter of chloroform was added to the reaction mixture as inhibitor and extractant of assay products. The reaction was measured against β-ionone control solutions under the same reaction conditions with cells carrying plasmid without insertion. Production was analyzed using GC and GC-MS.
Expression and purification of the optimization of recombinant DBR1
For protein expression, 1 ml of an overnight culture was used to inoculate 50 ml of LB kanamycin medium at 37 °C to an optical density at OD600 of 1.8. The expression of the proteins was induced by adding isopropyl-b-D-thiogalactoside (IPTG) to a final concentration of 0.6 mM, and the bacteria were further incubated at 27 °C for 18 h and gently shaken (180 rpm). Bacteria were harvested by centrifugation at 5000×g for 20 min at 4 °C, and then resuspended in 10 ml of 1× PBS.
Cells were lysed by sonification on ice with an MS 73 sonotrode (Bandelin Electronic, Berlin, and Germany) four times for 30 s at 10% of maximal power. Cell debris was removed by centrifugation (20,000 g, 30 min, and 4 °C). The recombinant protein was purified with Ni-chelating affinity chromatography(Novagen). The recombinant protein was eluted with an elution buffer containing increasing concentrations (20, 30, 50, 75, 100, 150, 200, and 300 mM) of imidazole. The protein was examined by SDS-PAGE. The protein concentration was determined by the Bradford method using BSA as a standard. The Bradford protein Assay Kit (Sangon Biotech, Shanghai, China) was employed for the determination.
The enzymatic activity of the recombinant DBR1 enzyme was assayed according to the method by Muangphrom et al.  and modified. The activity was determined GC using β-ionone as substrate. Assays contained 0.05 M TRIS-HCl, pH 6.5, 1 mM NADPH, 2 mM dithiothreitol (DTT), 1 mM β-ionone and 300 μg of the crude recombinant protein in a total volume of 500 μl at 45 °C for 10 min. After the incubation, 1 ml of chloroform was added to the reaction mixture as enzyme inhibitor and extractant of assay products. The enzymatic reaction was measured against β-ionone control solutions under the same reaction conditions while proteins were heat deactivated (100 °C, 10 min) prior to the incubation experiments.
The optimum temperature for enzyme activity was determined by standard assays ranging from 20 to 50 °C in the 0.05 M TRIS-HCl at pH 7.5. The optimum pH for DBR1 activity was determined by incubation at 45 °C for 10 min in a 0.05 M TRIS-HCl from pH 5.0 to 9.0. The results were expressed as percentages of the activity obtained at either the optimum pH or the optimum temperature.
To determine the effect of temperature on the stability of DBR1, the enzyme was pre-incubated in the 0.05 M TRIS-HCl (pH 6.5) for various times at 40, 45 and 50 °C in the absence of the substrate and other cofactor. The activity of the enzyme without pre-incubation was defined as 100%. The pH stability of the enzyme was determined by measuring the remaining activity after incubating the enzyme at 40 °C for 1 h in the 0.05 M TRIS-HCl from pH 5.0–9.0. Then, the residual activity of the enzyme incubated at variant pH was determined, immediately.
The effects of metals and chemical agents on the pure DBR1 enzyme were determined. Fe3+, Ca2+, Na+, Li+, K+, Mg2+, Zn2+, Al3+, Fe2+, NH4+, Mn2+, Cu2+, Ba2+, Hg2+, Co2+, Sr2+, Fe2+, DTT, and NADPH were assayed at the final concentrations of 1 and 5 mM in the reaction mixture. The control reaction was measured against under the same reaction conditions while DTT was absence. The effects of organic solvents on the enzyme were determined by adding 1, 2, 3, 4, 5 and 6% organic solvents (ethanol, methanol, and DMSO) to the reaction mixture.
The kinetic constant of DBR1 was determined by measuring the initial rates at various β-ionone concentrations (0.2, 0.4, 0.6, 0.8, 1.0, 1.2, 1.4, 1.6, and 1.8 mM). The reaction conditions were activated by 15 μg purified enzyme protein under standard conditions.
Analysis of GC and GC-MS
Dihydro-β-ionone was analyzed using GC 6890 system (Agilent, USA) and a HP5 column (30 m × 0.25 mm × 0.25 μm) with flame ionization detector. Injection and detector temperatures was 250 °C, Helium is employed as the carrier gas with the flow rate at 1 ml/min. Sample size and flow were 0.2 μl and 2 ml/min, respectively. Temperature-rising program was 50 °C for 2 min, raising to 120 °C at 10 °C/min for 2 min. Then the temperature was raised to 200 °C at 5 °C/min for 5 min. Dihydro-β-ionone was also verified using Trace ISQ-LT GC-MS (Thermo Fisher, USA) and a DB-5MS column (30 m × 0.25 mm × 0.25 μm) with flame ionization detector. Interface temperature was 250 °C, EI-MS with 70 EV ionization energy and swath range were 50–450 m/z.
Analyzing the sequence of four cDNA clones
Selection and production of dihydro-β-ionone on whole-cell systems
Expression and purification of the optimization of recombinant DBR1
Purification scheme for the recombinant protein
Ni affinity chromatography
Characterization of recombinant DBR1
Effects of metal cations and reagents on the recombinant enzyme activity
Cation of reagent
Cation of reagent
Effects of organic solvent on the recombinant enzyme activity
The kinetic parameters of the enzyme were analyzed using β-ionone as a substrate, the K M and Vmax values were 0.55 mM and 14.77 U/mg, respectively. DBR1 catalyzed the reduction of 2,3-unsaturated aldehydes, such as 2E-hexenal, 2E-nonenal, and artemisinic aldehyde. In the present investigation, although DBR1 showed a 10-fold higher specificity for 2E-nonenal relative to artemisinic aldehyde , specificity of similar strength was also shown for β-ionone relative to 2E-nonenal.
Analysis of dihydro-β-ionone via biological catalytic hydrogenation of β-ionone
In the modern synthetic biological study, the utilization of Gordian technique to invert β-ionone to dihydro-β-ionone needs the presence of an effective invertase. This task has rarely been reported. However, few publications are available regarding the conversion of unsaturated aldehydes and ketones as well as their genes. A clear distinction of substrate structure, could strongly affect enzymatic property and conversion efficiency. The enzyme employed in synthetic biology or biological catalytic conversion processes could break the boundaries of species. It could origin from the same or different plants or even may be rooted in microbes.
The artemisinin biosynthetic pathway has been studied for many years, and most of the enzymes have already been cloned [37–39]. In the artemisinin biosynthesis, artemisinic aldehyde Δ11(13) reductase (DBR) can convert artemisinic aldehyde into dihydroartemisinic aldehyde . DBR1 has been cloned by Zhang , which can perform catalytic hydrogenation of 2,3 unsaturated aldehyde. Artemisia annua aldehyde and hexene aldehyde can be catalyzed by DBR1. DBR2 as cloned by Zhang  which has high catalytic activity to cyclohexenone and carvone. OYE homologous genes i.e., Bac-OYE1 was cloned from Bacillus also exhibited activities toward α,β-unsaturated aldehydes, ketones, and other α,β-unsaturated compounds . In the present research, those four genetically engineered bacteria were screened to biotransform β-ionone to dihydro-β-ionone for the first time.
Dihydro-β-ionone is the characteristic aroma compounds of O. fragrans. There exists an enzyme in plants belonging to the genus Osmanthus Lour. (Fam.: Oleaceae) which could convert β-ionone into dihydro-β-ionone. Enzyme of plants might not be much effective, but it could be increasingly improved through repeated screening and research. Ro  published a design to assemble some genes from different organisms in order to establish non-natural metabolic pathways on yeast to synthesize largely precursor of artemisinin-artemisinic acid. This experiment simulated the natural plant’s metabolic process, to invert β-ionone to dihydro-β-ionone in vitro. Meanwhile, the present research team firstly discovered a kind of double bond reductase gene from non-O. fragrans, which has a strong transfer ability. Divalent cations were not necessary than monovalent cations for enzymatic activity and that DBR1 is not a metalloprotein. The DBR1 activity was not affected by DTT, which is a well-known thiol group inhibitor, suggesting that sulfhydryl groups may not be involved in the catalytic center of the protein structure. In addition, DBR1 was more stable at an acidic pH or a temperature range of 35–50 °C withβ-ionone, the cloning and characterization of Dbr1 present some biotechnological possibilities.
Based on this, in the present investigation, a recombinant strain pET-28a-DBR1 with steady catalytic performance through gene biotechnology has been achieved. This strain and its enzyme has commendable catalytic hydrogenation capacity to β-ionone’s exocyclic double bond.
Of the known plants, 2-alkenal reductases substrate specificity is somewhat broad, but without a complete substrate overlap among various enzymes. In the present study, four genetically engineered bacteria were screened to biotransform β-ionone to dihydro-β-ionone and only the DBR1 from A. annua showed high conversion efficiency. DBR1 was also expressed, purified, and biochemically characterized. It has a high catalytic efficiency for biotransforming β-ionone to dihydro-β-ionone, and thus becomes economically more feasible. This study, therefore, demonstrates that recombinant DBR1 has great potential for industrial applications, including bioconversion for producing natural compounds.
This study was funded by the Special Fund for Forest Scientific Research in the Public Welfare (Grant number 201404601), Jiangsu Key Lab of Biomass-based Green Fuels and Chemicals (Grant number JSBGFC14014), the 11th Six Talents Peak Project of Jiangsu Province (Grant number 2014-JY-011), and Priority Academic Program Development of Jiangsu Higher Education Institutions (PAPD).
Availability of data and materials
All data generated/analysed during the current study that are not already included in this published article, are available from the corresponding author on reasonable request.
LGZ and XSZ conceived and designed the experiments. XSZ and JJP analysed the data. XSZ wrote the first draft of the manuscript. SYL contributed to the writing of the manuscript. LGZ and FLC agree with manuscript results and conclusions. FT jointly developed the structure and arguments for the paper. LGZ and FLC made critical revisions and approved final version. All authors reviewed and approved of the final manuscript.
Ethics approval and consent to participate
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Baldermann S, Kato M, Kurosawa M, Kurobayashi Y, Fujita A, Fleischmann P, Watanabe N. Functional characterization of a carotenoid cleavage dioxygenase 1 and its relation to the carotenoid accumulation and volatile emission during the floral development of Osmanthus fragrans Lour. J Exp Bot. 2010;61(11):2967–77.View ArticleGoogle Scholar
- Liu CH, Long LP, Hou XB. Selective reduction of β-ionone during ultrasounic irridiation. J Cent South Univ. 2011;42(1):33–7.Google Scholar
- Beekwilder J, van Rossum HM, Koopman F, Sonntag F, Buchhaupt M, Schrader J, Hall RD, Bosch D, Pronk JT, van Maris AJ. Polycistronic expression of a β-carotene biosynthetic pathway in Saccharomyces cerevisiae coupled to β-ionone production. J Biotechnol. 2014;192(partB):383–92.View ArticleGoogle Scholar
- Ly MH, Hoang LC, Belin JM, Waché Y. Improved co-oxidation of beta-carotene to beta-ionone using xanthine oxidase-generated reactive oxygen species in a multiphasic system. Biotechnol J. 2008;3(2):220–5.View ArticleGoogle Scholar
- Rodríguez-Bustamante E, Sánchez S. Microbial production of C13-norisoprenoids and other aroma compounds via carotenoid cleavage. Crit Rev Microbiol. 2007;33(3):211–30.View ArticleGoogle Scholar
- Knowles WS. Asymmetric hydrogenations (Nobel lecture). ChemInform. 2002;33(40):1999–2007.Google Scholar
- Noyori R. Asymmetric catalysis: science and opportunities (Nobel lecture). Angew Chem. 2002;41(12):2008.View ArticleGoogle Scholar
- Richter N, Groger H, Hummel W. Asymmetric reduction of activated alkenes using an enoate reductase from Gluconobacter oxydans. Appl Microbiol Biot. 2011;89(1):79–89.View ArticleGoogle Scholar
- Belin JM, Bensoussan M, Serrano-Carreon L. Microbial biosynthesis for the production of food flavours. Trends Food Sci Technol. 1992;3:11–4.View ArticleGoogle Scholar
- Berger RG. Biotechnology of flavours—the next generation. Biotechnol Lett. 2009;31(11):1651–9.View ArticleGoogle Scholar
- Keasling JD. Synthetic biology and the development of tools for metabolic engineering. Metab Eng. 2012;14(3):189–95.View ArticleGoogle Scholar
- López J, Essus K, Kim IK, Pereira R, Herzog J, Siewers V, Nielsen J, Agosin E. Production of β-ionone by combined expression of carotenogenic and plant CCD1 genes in Saccharomyces cerevisiae. Microb Cell Factories. 2015;14(1):84.View ArticleGoogle Scholar
- Zhao YP, Mu XQ, Xu Y. Improvement in γ -decalactone production by Yarrowia sp. after genome shuffling. Chem Pap. 2014;68(8):1030–40.View ArticleGoogle Scholar
- Guo Y, Song H, Wang Z, Ding Y. Expression of POX2 gene and disruption of POX3 genes in the industrial Yarrowia lipolytica on the γ-decalactone production. Microbiol Res. 2012;167(4):246–52.View ArticleGoogle Scholar
- Etschmann MMW, Sell D, Schrader J. Medium optimization for the production of the aroma compound 2-phenylethanol using a genetic algorithm. J Mol Cata B Enzym. 2004;29(1–6):187–93.View ArticleGoogle Scholar
- Hua D, Ma C, Song L, Lin S, Zhang Z, Deng Z, Xu P. Enhanced vanillin production from ferulic acid using adsorbent resin. Appl Microbiol Biot. 2007;74(4):783–90.View ArticleGoogle Scholar
- Li YH, Sun ZH, Zhao LQ, Yan X. Bioconversion of isoeugenol into vanillin by crude enzyme extracted from soybean. Appl Biochem Biotech. 2005;125(1):1–10.View ArticleGoogle Scholar
- Pei XQ, Xu MY, Wu ZL. Two “classical” old yellow enzymes from Chryseobacterium sp. CA49: broad substrate specificity of Chr -OYE1 and limited activity of Chr -OYE2. J Mol Catal B Enzym. 2015;123:91–9.View ArticleGoogle Scholar
- Chaparro-Riggers JF, Rogers TA, Vazquez-Figueroa E, Polizzi KM, Bommarius AS. Comparison of three enoate reductases and their potential use for biotransformations. Adv Synth Catal. 2010;349(8–9):1521–31.Google Scholar
- Hall M, Stueckler C, Kroutil W, Macheroux P, Faber K. Asymmetric bioreduction of activated alkenes using cloned 12-oxophytodienoate reductase isoenzymes OPR-1 and OPR-3 from Lycopersicon esculentum (tomato): a striking change of stereoselectivity. Angew Chem. 2007;119(21):4008–11.View ArticleGoogle Scholar
- Kosjek B, Fleitz FJ, Dormer PG, Kuethe JT, Devine PN. Asymmetric bioreduction of α,β-unsaturated nitriles and ketones. Tetrahedron Asymmetry. 2008;19(12):1403–6.View ArticleGoogle Scholar
- Galili G, Galili S, Lewinsohn E, Tadmor Y. Genetic, molecular and genomic approaches to improve the value of plant foods and feeds. Crit Rev Plant Sci. 2002;21:167–204.View ArticleGoogle Scholar
- Burda E, Kraußer M, Fischer G, Hummel W, Müller-Uri F, Kreis W, Gröger H. Recombinant Δ4,5-steroid 5 β-reductases as biocatalysts for the reduction of activated C=C-double bonds in monocyclic and acyclic molecules. Adv Synth Catal. 2010;351(17):2787–90.View ArticleGoogle Scholar
- Goretti M, Ponzoni C, Caselli E, Marchigiani E, Cramarossa MR, Turchetti B, Buzzini P, Forti L. Biotransformation of electron-poor alkenes by yeasts: asymmetric reduction of ( 4S )-(+)-carvone by yeast enoate reductases. Enzyme Microb Technol. 2009;45(6–7):463–8.View ArticleGoogle Scholar
- Bruno Kilunga Kubata ZK, Nozaki T, Munday CJ, Fukuzumi S, Ohkubo K, Lazarus M, Maruyama T, Martin SK, Duszenko M, Urade Y. A key role for old yellow enzyme in the metabolism of drugs by Trypanosoma cruzi. J Exp Med. 2002;196(9):1241–51.View ArticleGoogle Scholar
- Frick UB. Characterization and cDNA-microarray expression analysis of 12-oxophytodienoate reductases reveals differential roles for octadecanoid biosynthesis in the local versus the systemic wound response. Plant J. 2002;32(4):585–601.View ArticleGoogle Scholar
- Me MC, Davis EM, Rushing GW, Croteau R. Monoterpene double-bond reductases of the (−)-menthol biosynthetic pathway: isolation and characterization of cDNAs encoding (−)-isopiperitenone reductase and (+)-pulegone reductase of peppermint. Arch Biochem Biophys. 2003;418(1):80–92.View ArticleGoogle Scholar
- Zhang Y, Teoh KH, Reed DW, Maes L, Goossens A, Olson DJ, Ross AR, Covello PS. The molecular cloning of artemisinic aldehyde Delta11(13) reductase and its role in glandular trichome-dependent biosynthesis of artemisinin in Artemisia annua. J Biol Chem. 2008;283(31):21501–8.View ArticleGoogle Scholar
- Richter N, Gröger H, Hummel W. Asymmetric reduction of activated alkenes using an enoate reductase from Gluconobacter oxydans. Appl Microbiol Biot. 2011;89(1):79.View ArticleGoogle Scholar
- Muangphrom P, Suzuki M, Seki H, Fukushima EO, Muranaka T. Functional analysis of orthologous artemisinic aldehyde Δ11(13)-reductase reveals potential artemisinin-producing activity in non-artemisinin-producing Artemisia absinthium. Plant Biotechnol. 2014;31(5):483–91.View ArticleGoogle Scholar
- Pereira PJ, Macedo-Ribeiro S, Párraga A, Pérez-Luque R, Cunningham O, Darcy K, Mantle TJ, Coll M. Structure of human biliverdin IXbeta reductase, an early fetal bilirubin IXbeta producing enzyme. Nat Struct Biol. 2001;8(3):215.View ArticleGoogle Scholar
- Xie J, Zhao D, Zhao L, Pei J, Xiao W, Ding G, Wang Z. Overexpression and characterization of a Ca2+ activated thermostable β-glucosidase with high ginsenoside Rb1 to ginsenoside 20(S)-Rg3 bioconversion productivity. J Ind Microbiol. 2015;42(6):839–50.Google Scholar
- Zhang YS, Teoh KHTH, Reed DWRW, Covello PSCS. Molecular cloning and characterization of Dbr1, a 2-alkenal reductase from Artemisia annua. Botany-Botanique. 2009;87(6):643–9.View ArticleGoogle Scholar
- Zhang X, Pei J, Zhao L, Tang F, Fang X, Xie J. Overexpression and characterization of CCD4 from Osmanthus fragrans and β-ionone biosynthesis from β-carotene in vitro. J Mol Catal B Enzym. 2016;134:105–14.View ArticleGoogle Scholar
- Youn B, Kim SJ, Moinuddin SG, Lee C, Bedgar DL, Harper AR, Davin LB, Lewis NG, Kang C. Mechanistic and structural studies of apoform, binary, and ternary complexes of the Arabidopsis alkenal double bond reductase At5g16970. J Biol Chem. 2006;281(52):40076–88.View ArticleGoogle Scholar
- Sun W. A kind of reducing coenzyme complex stabilizer. China Patent. CN101140279A. 2008.Google Scholar
- Arsenault PR, Vail D, Wobbe KK, Erickson K, Weathers PJ. Reproductive development modulates gene expression and metabolite levels with possible feedback inhibition of artemisinin in Artemisia annua. Plant Physiol. 2010;154(2):958–68.View ArticleGoogle Scholar
- Covello PS, Teoh KH, Polichuk DR, Reed DW, Nowak G. Functional genomics and the biosynthesis of artemisinin. Phytochemistry. 2007;68(14):1864–71.View ArticleGoogle Scholar
- Jiang W, Lu X, Qiu B, Zhang F, Shen Q, Lv Z, Fu X, Yan T, Gao E, Zhu M. Molecular cloning and characterization of a Trichome-specific promoter of Artemisinic aldehyde Δ11(13) reductase (DBR2) in Artemisia annua. Plant Mol Biol Rep. 2014;32(1):82–91.View ArticleGoogle Scholar
- Zhang H, Gao X, Ren J, Feng J, Zhang T, Wu Q, Zhu D. Enzymatic hydrogenation of diverse activated alkenes. Identification of two Bacillus old yellow enzymes with broad substrate profiles. J Mol Catal B Enzym. 2014;105(7):118–25.View ArticleGoogle Scholar
- Ro DK, Paradise EM, Ouellet M, Fisher KJ, Newman KL, Ndungu JM, Ho KA, Eachus RA, Ham TS, Kirby J, et al. Production of the antimalarial drug precursor artemisinic acid in engineered yeast. Nature. 2006;440(7086):940–3.View ArticleGoogle Scholar