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Reconstructing Biosynthetic Pathway of the Plant-Derived Cancer Chemopreventive-Precursor Glucoraphanin in Escherichia coli Han Yang, Feixia Liu, Yin Li, and Bo Yu ACS Synth. Biol., Just Accepted Manuscript • DOI: 10.1021/acssynbio.7b00256 • Publication Date (Web): 17 Nov 2017 Downloaded from http://pubs.acs.org on November 19, 2017

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ACS Synthetic Biology

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Revision of sb-2017-00256v.R2

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Reconstructing Biosynthetic Pathway of the Plant-Derived Cancer

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Chemopreventive-Precursor Glucoraphanin in Escherichia coli

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Han Yanga,b E-mail: [email protected]

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Feixia Liua E-mail: [email protected]

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Yin Lia,* E-mail: [email protected]

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Bo Yua,* E-mail: [email protected]

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Affiliations:

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a

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of Microbiology, Chinese Academy of Sciences, Beijing 100101, China

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b

CAS Key Laboratory of Microbial Physiological and Metabolic Engineering, Institute

University of Chinese Academy of Sciences, Beijing 100049, China

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* Corresponding authors.

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Phone/Fax: +86-10-64806132

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ACS Paragon Plus Environment


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Abstract

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Epidemiological data confirmed a strong correlation between regular consumption of

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cruciferous vegetables and lower cancer risk. This cancer preventive property is mainly

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attributed to the glucosinolate products, such as glucoraphanin found in broccoli that is

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derived from methionine. Here we report the first successful reconstruction of the

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complete biosynthetic pathway of glucoraphanin from methionine in Escherichia coli

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via gene selection, pathway design, and protein engineering. We used branched-chain

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amino transferase 3 to catalyze two transamination steps to ensure the purity of

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precursor molecules and used cysteine as a sulfur donor to simplify the synthesis

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pathway. Two chimeric cytochrome P450 enzymes were engineered and expressed in

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E. coli functionally. The original plant C-S lyase was replaced by the Neurospora

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crassa hercynylcysteine sulfoxide lyase. Other pathway enzymes were successfully

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mined from Arabidopsis thaliana, Brassica rapa, and Brassica oleracea. Biosynthesis

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of glucoraphanin upon co-expression of the optimized enzymes in vivo was confirmed

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by liquid chromatography-tandem mass spectrometry analysis. No other glucosinolate

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analogs (except for glucoiberin) were identified that could facilitate the downstream

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purification processes. Production of glucoraphanin in this study laid the foundation for

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microbial production of such health-beneficial glucosinolates in a large-scale.

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Keywords: glucoraphanin, pathway engineering, protein engineering, microbial

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synthesis

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2


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Abstract Graphic

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Plants can produce a wide range of secondary metabolites, many of which are

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valuable pharmaceutical and nutraceutical compounds.1 Chemical synthesis of these

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secondary metabolites have been a challenge due to the complexity of their structures.

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Consequently, it is more favorable to obtain them by isolation from plants or

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semisynthesis.2,3 However, only small quantities of such compounds are generated in

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their host plants at particular time points during development, which in turn may make

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it difficult for large-scale production. Over the past two decades, it has become

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technically feasible to introduce the natural synthetic pathways from plants to

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microorganisms for production purposes, thus bypassing the above limitations. Some

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notable examples include the antimalarial compound artemisinic acid, the antioxidant

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resveratrol, the flavor component vanillin, the universal sesquiterpene precursor,

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farnesyl diphosphate, and taxadiene (a precursor of the anticancer agent taxol).1

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Glucosinolates are naturally produced by members of cruciferous vegetables and

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contribute to the health-beneficial effects.4 In particular, glucoraphanin (GRA), a

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glucosinolate common among broccoli and Arabidopsis thaliana is associated with

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reduced risks for cardiovascular diseases and cancer.5 Conversion of GRA upon

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consumption could be mediated by myrosinases from endogenous plant or gut

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microorganisms, resulting in the formation of sulforaphane (SFN).6 Studies performed

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with animals as well as cellular models have indicated that SFN possesses

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cancer-preventive properties, such as antioxidation, anti-inflammation, induction of

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apoptosis and cell-cycle arrest.7 Very recent research has confirmed that SFN can also

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improve glucose control in type-II diabetes patients.8 These findings have drawn

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considerable attention to the optimization of GRA production for pharmaceutical

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applications, as well as for use as a dietary supplement.

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Glucosinolate biosynthesis, which has been extensively studied in Arabidopsis, 4


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comprises up to three stages, namely (i) side chain elongation, (ii) core structure

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formation, and (iii) secondary modifications.1 The three independent stages of the

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GRA-biosynthesis pathway in plants are shown in Figure 1. First, methionine

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undergoes a string of reactions that mediate one or more rounds of chain elongation,

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reminiscent of the conversion from valine to leucine. Elongation of methionine with

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two methyl groups leads to the formation of dihomo-methionine (DHM), the precursor

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of GRA. The initial cytosolic transamination process is accomplished by

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branched-chain amino transferase 4 (BCAT4), resulting in the α-keto acid (αKA).9 The

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αKA

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methylthioalkylmalate

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2-(2-methylthio) ethylmalate derivative, which subsequently undergoes isomerization

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and decarboxylation. To be specific, the ethylmalate derivative is catalyzed by an

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isopropylmalate isomerase (IPMI), as well as an isopropylmalate dehydrogenase

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(IPMDH) to form a chain-elongated αKA.10-12 Two cycles of chain-elongated αKA

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(2-oxo-6-methylthiohexanoic acid) is further transaminated by BCAT3 to form DHM.

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Later, DHM enters the second biosynthetic stage where the corresponding

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glucosinolates were formed. The pathway responsible for core structure formation

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involves the activities of five cytosolic enzymes. They include cytochrome P450

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enzymes from the CYP79 and CYP83 families that mediate two monooxygenation

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reactions of forming corresponding oximes, and then to reactive compounds.4

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S-alkylthiohydroxamates are formed in a sulfurization step, which may involve either a

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non-enzymatic step or an enzyme similar to glutathione-S-transferase.13 The other three

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cytosolic enzymes are responsible for the last three enzymatic steps: (i) a C-S lyase,

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converting S-alkylthiohydroxamates to thiohydroxamates,14 (ii) a glucosyltransferase,

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transferring glucose to thiohydroxamates to form desulfoglucosinolates,15 and (iii) a

is

transported

into

chloroplasts

synthase

(MAM)

where with

it

is

condensed

acetyl-CoA

5


to

by

produce

the a


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sulfotransferase, adding a sulfate group to produce glucosinolates.16,17 Finally, the

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product is modified by a flavin monooxygenase to generate GRA.18

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Although it is feasible to transfer the biosynthetic pathways from plants to

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microorganisms for production, exactly how to express the proteins functionally is the

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first challenge to overcome. Some enzymes derived from plants are difficult to express

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functionally in prokaryotes because they undergo different post-translational

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modifications and adopt structures distinct from that generated in eukaryotic cells.

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Previous studies have shown that many efforts are needed to optimize the heterologous

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production of eukaryotic proteins in microorganisms. For example, expression of the

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hydrophobic membrane protein P450s in E. coli is especially challenging due to its

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tendency to form insoluble inclusion bodies.19 Moreover, some enzymes, such as the

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C-S lyase SUR1 in Arabidopsis, whose function is not redundant, have been found to

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be very unstable in E. coli.14 Thus, exploring isoenzymes from different sources is

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needed. In addition, plants produce over 100 glucosinolates and these compounds are

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usually mixed together owing to the multifunctional and redundant nature of the

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enzymes, making it necessary to select relatively efficient enzymes and design a

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pathway to reduce by-product assembly in microorganisms. For example, in the first

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stage of GRA production, methionine is converted to DHM often with the by-products

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of homo-methionine (HM), trihomo-methionine, and other derivatives. Establishing

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how to shift the flux towards maximum DHM production is also highly desirable.

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In this study, we first selected appropriate enzymes and functionally expressed all

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necessary enzymes for GRA production in E. coli. Then, we engineered the

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biosynthetic pathway from methionine to GRA, which was transferred from plants to

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microbes. After introducing 10 enzymes from different sources, the final product GRA

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was confirmed by LC-MS/MS analysis. Our results suggest that heterologous 6


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production of glucoraphanin in E. coli is a promising way to generate this valuable

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compound.

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Results and Discussion

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Selection and expression of enzymes in the side chain-elongation pathway. The

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side chain-elongation pathway involves several transamination steps.20 The initial

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transamination step is catalyzed by BCAT4,9 while BCAT3 mediates the terminal

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chain-elongation steps and can also partially replace the function of BCAT4.21

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Moreover, BCAT3 only participates in the extension of methionine by adding one or

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two methyl groups. The enzyme converts 5-methylthio-2-oxopentanoate to HM and

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6-methylthio-2-oxohexanoate to DHM, whereas less enzymatic activity is measured for

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the conversion of 4-methylthio-2-oxohexanoate to trihomo-methionine.21 To reduce the

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formation of byproducts and to simplify the pathway, we selected one branched amino

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transferase (BCAT3) for the side chain-elongation pathway since only DHM is a

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precursor for glucoraphanin production. By deleting the potential chloroplast-targeting

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signal (60 amino acids), the truncated BCAT3 was solubly expressed in E. coli

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(Supplementary file, Fig. S1a).

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Glucosinolates in Arabidopsis normally contain a variable side chain with additional

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one to six methyl groups as a result of chain elongation of methionine derivatives.22

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GSL-ELONG is one of the critical loci responsible for such variation. It directs the

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degree of side chain elongation of methionine-derived glucosinolates.23 Previous

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cloning of the GS-ELONG QTL gene has led to the identification of the MAM1,

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MAM2, and MAM3 genes.24,25 MAM2 plays an important role in producing aliphatic

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glucosinolates by adding one methyl group, while MAM1 mediates the production of

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glucosinolates up to two elongation cycles,24-26 and MAM3 is involved in all aliphatic 7



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glucosinolates production but prefers to add one, five and six methyl groups,

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respectively.26,27 After transferring the DHM biosynthetic pathway into E. coli with

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MAM1, the byproduct HM still represented a substantial portion of the total output,28

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which is not favorable for GRA production. Aliphatic glucosinolates derived from HM

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are termed “3C” glucosinolates, whereas those derived from DHM are called “4C”

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glucosinolates.24 In Brassica oleracea, Li et al. confirmed that the 3C and 4C

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glucosinolates are independently regulated by the GSL-PRO and GSL-ELONG genes.29

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Therefore, it is possible to channel the glucosinolate pathway with increased content of

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glucoraphanin by only optimizing the GSL-ELONG expression. Here, we selected

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GSL-ELONG (encoding methylthioalkylmalate synthase) from B. oleracea to control

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the condensation reaction for reduced byproducts. The truncated GSL-ELONG gene,

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without the signal sequence was successfully expressed in E. coli (Supplementary file,

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Fig. S1b).

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IPMI is a heterodimer with a large subunit (LSU) and a small subunit (SSU). In

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Arabidopsis, there are one LSU gene (IPMI LSU1) and three genes for SSU (IPMI

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SSU1, SSU2, and SSU3).9,30 While the large subunit is responsible for both leucine

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biosynthesis and methionine chain elongation,31 the three small subunits may specialize

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in different functions. IPMI SSU2 and IPMI SSU3 function in methionine chain

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elongation, while IPMI SSU1 is important in leucine biosynthesis.31,32 In Arabidopsis,

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three IPMDHs are crucial for methionine chain-elongation. IPMDH1 is mainly

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participated in glucosinolate biosynthesis, whereas both IPMDH2 and IPMDH3 also

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contribute to leucine biosynthesis.33-35 Mirza et al. introduced IPMI (LSU1 and SSU3

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subunits) and IPMDH1 into E. coli to successfully construct the DHM-biosynthesis

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pathway.28 Therefore, we also selected IPMI (LSU1 and SSU3) and IPMDH1 of

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Arabidopsis for this study. The N-terminal sequences predicted to be responsible for 8


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chloroplast localization of the three genes were deleted as previously described.32,33

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The soluble expression of the modified IPMI (LSU1 and SSU3) and IPMDH1 was

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confirmed (Supplementary file, Fig. S1c).

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As illustrated by literatures, several isozymes are involved in each step of the side

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chain-elongation pathway. These isozymes catalyze similar reactions with subtle

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difference for preferences in substrates. This phenomenon creates redundant expression,

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which may be necessary for plants, but it would have led to a heavy metabolic load for

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microorganisms if we introduce all relevant genes for DHM biosynthesis. Therefore, to

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reduce the number of by-products formed and lower the burden to microbial strains, we

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chose BCAT3 (derived from Brassica rapa), GSL-ELONG (derived from B. oleracea),

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IPMI (LSU1 and SSU3), and IPMDH1 (derived from Arabidopsis thaliana) to catalyze

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the first steps of glucoraphanin production from methionine to DHM.

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Selection and expression of enzymes in the core structure-formation pathway.

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CYP79 family cytochrome P450s mediate the conversion of amino acids to oximes

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during glucosinolate biosynthesis, whereas CYP83-family members are involved in

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subsequent oxime metabolism. When functioning together, the CYP79F1/F2 and

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CYP83A1 enzymes result in the production of aliphatic glucosinolates, while the

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CYP79A2/B2/B3 and CYP83B1 enzymes generate indolic/benzenic glucosinolates.

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Compared to CYP79F2, which yields exclusively long-chain aliphatic glucosinolates,

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the CYP79F1 enzyme can catalyze mono- to hexahomo-methionine.20 Considering the

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specificity of the enzymes, we chose CYP79F1 (B. oleracea) and CYP83A1 (B. rapa)

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to metabolize short-chain aliphatic amino acids and oximes.

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Application of the plant P450 enzymes in E. coli is restricted by two major factors,

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with the first one being the lack of cytochrome P450 reductases (CPRs), which are 9



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responsible for electron transfer in eukaryotes.36 The second one is the absence of

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compatible membrane-bound sequences.37,38 Modification at the N-terminal of P450s

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have resulted in successfully expression in prokaryotic bacteria.39 In this study, the

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modification procedures used for functionally expressing CYP79F1 and CYP83A1 in E.

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coli were performed as described in our previous report for expressing CYP79A2 and

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CYP83B1.40 Briefly, the first seven amino acids of CYP79F1 were replaced with a

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synthetic mammalian N-terminal peptide ε to create CYP79F1[ε:8-540].37 Then, the 3′

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terminus of the modified CYP79F1[ε:8-540] enzyme was joined to the 5′ terminus of

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ATR2 [73-711] using an artificial linker (λ) to create a fusion protein (Fig. 2a). The

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used λ fragment was chosen as GSTSSGSG.41 Sodium dodecyl sulfate-polyacrylamide

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gel electrophoresis (SDS-PAGE) analysis indicated the successful expression of the

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chimeric protein in E. coli (Fig. 2b).

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Cytochrome P450 enzyme CYP83A1 is the second enzyme in the core

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structure-biosynthesis pathway, converting aldoxime to an aci-nitro compound. We

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modified CYP83A1 in the same manner as done for CYP79F1. CYP83A1 contains two

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predicted transmembrane domains, one of which is in the N-terminal region and the

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other is in the central region of the primary sequence. These putative transmembrane

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domains increase the risk of failure for protein expression and proper folding.

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Fortunately, we successfully expressed the chimera enzyme by only modifying the

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N-terminal transmembrane sequence to generate a fusion protein comprised of

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CYP83A1[ε:20-501] with a truncated ATR2[73-711] domain (Fig. 2b).

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Our previous investigation has demonstrated that the product catalyzed by

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CYP83B1, nitrile oxide, can interact with cysteine spontaneously to generate

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S-alkylthiohydroxamates for downstream reaction catalyzed by the C-S lyase.40 Thus,

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in this study, we also used cysteine as sulfur donor in vivo to omit two enzymes 10


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(glutathione-S-transferase GSTF and γ-glutamyl peptidase GGP1). The three

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subsequent steps of glucoraphanin synthesis are mediated by the so-called

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post-aldoxime enzymes, which are low selective for the side chain but highly selective

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for the core group.42 SUR1 has been characterized as a C-S lyase in glucosinolate

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biosynthesis and the function of SUR1 is not redundant in plants.14 Since the

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recombinant SUR1 protein is quite unstable in heterologous host14, we tested enzymes

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with similar functions from other sources. In another sulfur-containing molecule in the

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ergothioneine-biosynthetic pathway of the fungus Neurospora crassa, a novel C-S

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lyase hercynylcysteine sulfoxide lyase (EGT2) was identified and in vitro

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reconstitution revealed that it catalyzes a similar reaction with that of SUR1 (Fig. 3).43

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Thus, this enzyme attracted our attention and the corresponding gene was synthesized

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after codon optimization and successfully expressed in E. coli in its active form (Fig.

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3).

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The next-to last step in the core structure-formation pathway is glucosylation, which

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catalyzed by a UDP-glucose-thiohydroximate glucosytransferase. UGT74B1 is the

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major enzyme catalyzing this reaction in vivo.15 The codon optimized UGT74B1 was

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easily expressed in E. coli, as previously described40 and is shown in Figure 3.

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Desulfoglucosinolate: PAPS sulfotransferase catalyzes the sulfurization of

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desulfoglucosinolates during the final step of core structure formation in glucosinolate

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biosynthesis. Three sulfotransferases with different substrate preferences are present in

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plants, classified as ST5a, ST5b, and ST5c. ST5a prefers tryptophan- and

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phenylalanine-derived desulfoglucosinolates, while aliphatic desulfoglucosinolates

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serve as the preferred substrates of ST5b and ST5c.16,17 Moldrup et al. transformed the

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genes involved in benzyl glucosinolate synthesis in tobacco plants and successfully

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enabled them to synthesize benzyl glucosinolate.44 This finding showed that the supply 11



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of the co-substrate PAPS was a bottleneck in synthesis, rather than the sulfotransferases.

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Because the binding affinity of the S donor PAPS is higher for ST5c,17 we chose ST5c

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as the candidate enzyme in this study and the codon-optimized ST5c gene was

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successfully expressed in E. coli without other attempts (Fig. 3).

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Selection and expression of enzymes in secondary modification. Hanse et al. have

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identified the flavin monooxygenase (FMO) enzyme in Arabidopsis, FMOGS-OX1,

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which converts methylthioalkyl glucosinolates to methylsulfinylalkyl glucosinolates.18

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FMOGS-OX1 efficiently converted 4-methylthiobutyl glucosinolate (derived from

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dihomo-methionine) to glucoraphanin, but only about 60% of 3-methylthiopropyl

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glucosinolate

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3-methylsulfinylpropyl glucosinolate.6 Thus, we chose FMOGS-OX1 from Arabidopsis

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as a suitable FMO to modify the side chain in this study. We directly expressed the gene

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amplified from the complementary DNA (cDNA) of Arabidopsis without any changes

260

in the gene sequence (Fig. 3).

(derived

from

homo-methionine)

was

converted

to

261 262

Functional verification of core formation and side chain-modification pathways

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in vitro. After successfully expressing all the pathway enzymes in E. coli, their

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enzymatic activities needed to be verified. Because most intermediates were not

265

commercially available and were difficult to detect, it was challenging to measure the

266

activity of each enzyme. Given these limitations, we opted to use a combination of pure

267

enzymes to detect product formation and verify their enzymatic activities. The first part

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(side chain elongation) of glucoraphanin synthesis in E. coli was already achieved

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previously, although different sources of enzymes were selected for this study.28 In

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addition, the decomposition product of glucoraphanin (sulforaphane) was readily 12


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detected at low levels by GC-MS. Thus, we further added Brevicoryne brassicae

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myrosinase to the reaction system containing only the enzymes required for the core

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formation and side chain-modification pathways, in order to test the activities.40

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Reaction was performed by incubating all the enzymes, substrates and cofactors

275

involved at 25°C for 1 h. The sulfur donor GSH was replaced by cysteine for the

276

reaction.40

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GC-MS analysis showed that 3-butenyl isothiocyanate was detected in the reaction

278

system (Fig. 4). The presence of 3-butenyl isothiocyanate has been proposed as the

279

result of thermal degradation of sulforaphane during sample injection in GC or GC-MS

280

analysis.45 Blazevic et al. also reported that some 3-butenyl isothiocyanate could

281

originate from sulforaphane degradation. In our experiments, direct detection of the

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authentic standard sulforaphane by GC-MS also revealed the presence of 3-butenyl

283

isothiocyanate (Fig. 4c).46 In addition, since DHM was not commercially available, we

284

used HM as the substrate. LC-MS analysis of the substrate HM was also accompanied

285

by small amount of DHM, which could have originated as an impurity of fmoc

286

L-homo-methionine. We anticipated that iberin (derived from HM) would be the main

287

product with small amount of sulforaphane (derived from DHM), whereas only

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sulforaphane was identified in our studies. This outcome might have been due to the

289

enzyme specificities, although the speculation needs to be verified. Nevertheless,

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identification of 3-butenyl isothiocyanate indicated that sulforaphane was formed in the

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reaction and most importantly, confirmed the activities of all enzymes expressed in E.

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coli, which encouraged us to assemble the entire pathway in vivo.

293 294

Assembling the entire glucoraphanin-biosynthesis pathway in E. coli.

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Sulforaphane is known to be unstable upon exposure to heat and could be degraded into 13



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several volatile compounds.47,48 In addition, sulforaphane also has antimicrobial

297

activity. To achieve stable product formation by microbial fermentation, we introduced

298

the entire glucoraphanin-synthesis pathway in E. coli, starting from methionine. The

299

whole pathway (including the enzymes engineered in the current study), in comparison

300

with the natural pathway in Arabidopsis thaliana, is shown in red in Figure 5.

301

The enzymes mediating side chain elongation, core structure formation, as well as

302

side chain modification were co-expressed together. The synthetic operons were all

303

under the control of the T7 promoter (Supplementary file, Fig. S2). The first operon

304

consisted of the first two genes involved in the side chain-elongation biosynthetic

305

pathway, while the second operon, which was driven by the T7 promoter in a different

306

plasmid (pET-28a), comprised the remaining genes of the DHM-biosynthetic pathway

307

in

308

BCAT3-GSL-ELONG-IPMDH1-IPMI-LSU1-IPMI-SSU3. It is widely accepted that

309

higher expression is observed from the upstream genes in the operon than the genes put

310

at downstream.49 Therefore, we positioned IPMDH1 upstream to IPMI (LSU1 and

311

SSU3) in the second operon to increase the activity of IPMDH1. The four genes in the

312

transcriptional order of EGT2-UGT74B1-ST5c-FMOGS-OX1 were inserted into multiple

313

cloning site 1 (MCS1) of pACYCDuet-1, and the forth operon was composed of

314

CYP79F1-CYP83A1-ATR2 and inserted into MCS2. The optimum proportion of P450

315

enzyme to CPR was ∼15:1 in natural plant systems.50 This trend demonstrates the

316

chimera of P450 and CPR with a ratio of 1:1 was not the most effective scheme. High

317

expression of P450s consumed a huge amount of resource and resulted in heavy

318

metabolic burden, which in turn depressed productivity.51 Therefore, P450s catalyzed

319

two consecutive steps, which should be the rate-limiting steps, and then we fused the

320

modified CYP79F1 and CYP83A1 genes with one ATR2 gene in the final position of

the

transcriptional

14


order

of

Page 15 of 42

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321

the operon to maintain relatively low expression.

322 323

In vivo production of GRA in E. coli harboring the modified synthesis pathway.

324

The strain with two plasmids harboring four operons was constructed and analyzed for

325

synthesis of the final product glucoraphanin, derived from methionine. Because so

326

many transgenes were co-expressed in single cells, we first conducted RT-PCR

327

analysis to confirm that all the genes were successfully assembled in E. coli and that the

328

added artificial ribosome binding sequences in front of each gene were functional. All

329

genes in the glucoraphanin biosynthesis pathway were detectable by RT-PCR analysis

330

(Supplementary file, Figure S3), indicating that they were all transcribed correctly.

331

Next, we further investigated the product from methionine transformation in

332

isopropyl β-D-thiogalactopyranoside (IPTG)-induced cells. LC-MS/MS analysis

333

results showed that a trace amount of glucoraphanin was produced in the reaction

334

system (Fig. 6). As expected, glucoiberin was also detected as the only byproduct.

335

Glucoraphanin was detected both in the cells and medium, while glucoiberin was only

336

detected in the cells. The ratio of glucoraphanin/glucoiberin in the cells was about 3: 2.

337

The lack of detectable glucoiberin in the medium may have resulted from a low total

338

yield of glucosinolates and a negative matrix effect. Nonetheless, this finding indicated

339

that the reconstructed pathway in E. coli could not efficiently distinguish between “4C”

340

and “3C” glucosinolates and this result may have been due to low GSL-EONG

341

expression, which suggested a further target for optimizing biosynthesis at the pathway

342

level. The result was also in accordance with previous observations following transient

343

expression of glucoraphanin in tobacco plants,6 in which both glucoraphanin and

344

glucoiberin were detected. Although the native CYP79F1 enzyme purified from

345

Brassica can metabolize mono- methionine to hexahomo-methionine,20 glucosinolate 15



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

346

derivatives more than “4C” were not detected in our experiments. This could be due to

347

the selection of BCAT3 in the first stage pathway to avoid other byproducts, where the

348

substrate specificity of the enzyme only facilitated catalysis of two rounds of

349

condensation reactions at most. Furthermore, CYP79F1 exclusively catalyzes aliphatic

350

amino acid monooxygenase reactions, which could explain why no other

351

amino-acid-derived products were detected in this study. Thus, the enzyme-selection

352

strategy ensured the purity of the final product, which will reduce the number of

353

downstream purification steps required and the processing costs. The absence of

354

pathway intermediates identified in the spectrum could have been due to the relatively

355

low concentrations, where the intermediates were quickly transformed to the

356

downstream compounds. Alternatively, they simply could not be identified due to the

357

lack of authorized standards.

358

In summary, enzyme candidates from different sources were selected by reviewing

359

the literature. The N-terminal domains of two membrane P450 proteins were modified

360

for expression on the membrane of E. coli cells to facilitate protein expression. The

361

P450 proteins were also modified as fusion protein with the corresponding P450

362

reductases to achieve electron transfer. In addition, some enzymes, which could not be

363

expressed in soluble form or were not stable in E. coli cells, were substituted with

364

isoenzymes from different sources. By using cysteine as the sulfur donor in a

365

non-enzymatic step and one BCAT enzyme (BCAT3) to participate in both the

366

beginning and the final transamination reactions, we simplified the synthesis pathway,

367

avoided impurities, and lightened the metabolic burden to the cells. Our study

368

represents the first case where glucoraphanin was synthesized from methionine by

369

microbial cells. At present stage, the glucoraphanin levels in the broth were rather low

370

and the concentrations were roughly estimated to be 2-3 µg/L, despite of the 16


Page 16 of 42

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371

complicated sample-processing procedure limited the accuracy of such measurements.

372

Although some technical obstacles remain that may impede the process of large-scale

373

production of glucoraphanin by E. coli strains at present, the achievements in this study

374

provided a start for further improving the production performance. Importantly, the

375

protein-expression and gene-mining processes revealed by this study could also

376

provide reference information for demonstrating the biosynthesis of other secondary

377

plant products by microorganisms. More engineering strategies are required to increase

378

the efficiency, including further optimization of P450s expression. The changes such as

379

reduction of secondary mRNA structure, bacterial codon usage, the use of molecular

380

chaperones, as well as varying external growth conditions, also appear to influence

381

P450s expression in prokaryotic organisms.52 We are currently evaluating systems

382

metabolic engineering to increase the supplies of co-substrates, to construct ATP and

383

NADPH cofactor-regeneration cycles, and to balance the protein expression at the

384

pathway level to strengthen the production capacity of glucoraphanin.

385 386

Methods

387

Functional expression of individual enzymes in E. coli. The sequences of primers

388

used in this study for recombinant protein expression are shown in Table 1. The

389

BCAT3 gene from B. rapa and the GSL-ELONG gene from B. oleracea (encoding

390

methylthioalkylmalate synthase) were codon-optimized and synthesized by GENEWIZ

391

Co., Ltd. (Suzhou, China). IPMDH1, IPMI-LSU1, and IPMI-SSU3 were directly

392

amplified from Arabidopsis cDNA. The cytochrome P450 genes CYP79F1 from B.

393

oleracea and CYP83A1 from B. rapa, EGT2 (encoding a C-S lyase) from N. crassa,

394

UGT74B1 (encoding a glucosyltransferase) from B. rapa, and ST5c (encoding a

395

sulfotransferase) from B. rapa were codon-optimized and synthesized by GENEWIZ. 17



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

396

FMO-GSOX1 (encoding a flavin monooxygenase) was directly amplified from

397

Arabidopsis cDNA. All gene sequences used in this study can be accessed in the

398

associated online supplementary file.

399

The pET-28a and pETDuet-1 vectors were used for recombinant protein expression.

400

The chain-elongation genes BCAT3, GSL-ELONG, IPMI (IPMI-LSU1 and IPMI-SSU3

401

fusion genes), and IPMDH1 were constructed in pET-28a after truncating their

402

respective membrane-positioning sequences. For the core structure-formation pathway,

403

the CYP79F1, CYP83A1, EGT2, UGT74B1, and ST5c genes were constructed in the

404

pET-28a vector. CYP79F1 and CYP83A1 were each fused with a truncated variant of

405

the CPR gene ATR2 and expressed as chimeras after substituting their predicted

406

transmembrane domain segments with that of bovine 17α-hydroxylase, as described

407

previously with minor changes.40 The gene encoding the secondary-modification

408

enzyme FMO-GSOX1 was inserted into pETDuet-1. E. coli DH5α cells were used for

409

plasmid propagation, and BL21(DE3) cells were used for protein expression. The

410

molecular chaperone pGro7 was used for FMO-GSOX1 expression to enhance the

411

purification efficiency. Protein expression and purification were performed in

412

accordance with our previous work.40

413 414

Reconstitution of the core structure formation and side-chain modification

415

pathways in vitro. The individual genes were expressed, purified and added to the

416

reaction system for biosynthesizing sulforaphane, except for CYP79F1 and CYP83A1

417

since it is hard to get pure enzymes and only the respective cell lysates were used. Since

418

no commercial source of DHM was available, we used commercial fmoc

419

L-homo-methionine to produce HM after removing the fmoc-group.53 Synthesis of the

420

resultant HM molecule was verified by LC-MS analysis. The reaction buffer for 18


Page 18 of 42

Page 19 of 42

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421

sulforaphane biosynthesis contained 5 mM homo-methionine, 5 mM cysteine, 2 mM

422

UDP-glucose, 2 mM pyridoxal phosphate (PLP), 10 mM NADPH, and 2 mM

423

3'-phosphoadenosine-5'-phosphosulfate lithium salt (PAPS). The reaction was

424

performed at 25°C for 1.5 h. Next, an equal volume of dichloromethane was used to

425

extract the reaction mixture twice at room temperature. The collected dichloromethane

426

portion was evaporated and re-dissolved in 100 µl dichloromethane for subsequent

427

GC-MS analysis.

428 429

Integration of the glucoraphanin-synthesis pathway into E. coli. The pET-28a

430

and pACYCDuet-1 vectors were used for co-expressing all genes in the

431

glucoraphanin-synthesis pathway in clonal E. coli cells. The primer sequences used for

432

pathway reconstruction are shown in Table 2. The T7 promoter was used for sequential

433

expression of glucoraphanin-biosynthesis genes. The plasmid pET-28a harbored genes

434

driving the first step of glucoraphanin biosynthesis (chain elongation), including

435

BACT3, GSL-ELONG, IPMI (LSU1 and SSU3), and IPMDH1. The plasmid

436

pACYCDuet-1 was used to express genes driving core structure formation (CYP79F1

437

and CYP83A1 fused with ATR2, EGT2, UGT74B1, and ST5c) and modification

438

(FMOGS-OX1). The respective ribosome-binding site was put in front of each gene to

439

make the proper expression. The pET-28a and pACYCDuet-1 co-transformants were

440

picked up from LB agar plates containing 50 µg/ml ampicillin and 12.5 µg/ml

441

chloramphenicol, respectively.

442 443

Verification of gene expression by RT-PCR analysis. To check the expression of

444

genes encoding the whole glucoraphanin-synthesis pathway, total RNA isolation

445

followed by RT-PCR analysis was performed. Aliquots of 4-mL overnight-grown 19



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

446

cultures were subsequently centrifuged for 1 min at 13,000 × g. RNA was isolated using

447

the E.Z.N.A.TM Bacterial RNA Kit (Omega Bio-Tek, USA). Total RNA concentrations

448

were determined based on the absorbance at 260 nm (NanoVue, GE Healthcare).

449

cDNAs were prepared with the FastQuant RT Kit (with gDNase) (Tiangen, China),

450

using appropriate gene-specific primers.

451 452

Transforming methionine to GRA by resting cell reactions. A single

453

transformant was inoculated into 10 mL LB media with 50 µg/ml ampicillin and 12.5

454

µg/ml chloramphenicol and grown overnight. The cultures were diluted at 1:50 in 200

455

mL TB media containing antibiotics and additional 75 µg/ml δ-aminolevulinic acid was

456

added when the cells grew to an OD600 of ~0.2. When the OD600 further reached 0.6-0.8,

457

0.5 mM IPTG was added for an additional growth for 12 h at 37 ºC to promote the

458

protein expression. The cells were collected and re-suspended in M9 medium

459

supplemented with 5 mM methionine, 5 mM cysteine, 0.2 mM PLP, 0.2 mM PAPS,

460

and 0.1 mM Fe(II) at an OD600 of ~30. After cultivation at 30 ºC for 48 h, the

461

supernatant was collected after centrifugation and freeze-dried. Pure methanol was

462

added to dissolve the lyophilized powder, the resulting solution was put at room

463

temperature for 4 h, and the supernatant was collected and filtered through a 0.22-µm

464

polyvinylidene fluoride filter. Then, the supernatant was further evaporated and

465

re-dissolved in 100 µl deionized water for LC-MS/MS analysis.

466 467

Analytical methods. GC-MS was performed on a Shimadzu QP2010 Ultra gas

468

chromatograph coupled to a Shimadzu mass spectrometer. A DB-5 column (30 m ×

469

0.25 mm × 0.25-µm) was used (head pressure, 100-kPa, splitless injection). For

470

sulforaphane, the analysis conditions used were as follows: injection port 250 °C; the 20


Page 20 of 42

Page 21 of 42

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471

oven temperature program was held at 50 °C for 2 min, increased at 10 °C min-1 to

472

250 °C and held for 5 min; the ion source was run in EI mode (70 ev) at 200 °C.

473

Separation of glucosinolates were achieved by high-performance liquid

474

chromatography (HPLC) coupled with a Zorbax Eclipse Plus C18 column (250 mm ×

475

4.6 mm × 5 mm, Agilent Technologies, Germany). Formic acid (0.05%) in water and

476

methanol were employed as mobile phases A and B respectively. The injection volume

477

was 20 µL. The gradient elution program was applied at a flow rate of 0.5 mL/min as

478

follows: 0-30 min, 5% B in A; 30-32 min, 5-100% B in A; 32-40 min 100% B, 40-41

479

min 100-5% B in A and 41-50 min 5% B. The column temperature was maintained at

480

25 °C. The Agilent 1260 HPLC system was coupled to an AB SCIEX QTRAP 4500

481

mass spectrometer (Foster, CA, USA) equipped with an electrospray ion source

482

operated in negative ionization mode. The operating parameters were as follows:

483

collision gas: 20.0; collision gas: medium; ion spray voltage: -4,500 V; Temperature:

484

500 °C; ion source gas 1: 60.0; ion source gas 2: 60.0; declustering potential: -82.0;

485

entrance potential: -8.0; collision energy: -40.0; collision cell exit potential: -18.0.

486

Glucoraphanin ion detection was performed by multiple reaction monitoring (MRM)

487

mode at m/z 436-96.8, 436-177.8, 436-259.0 and 436-371.9. The same operating

488

parameters were used to detect glucoiberin, except that the collision energy was -26.0.

489

Ion detection was performed in MRM mode at m/z 422-96.9, 422-195.8, 422-258.9 and

490

422-357.9, according to previous literature.9

491

492

Supporting Information

493

Soluble enzyme expression data for BCAT3, GSL-ELONG, IPMI (LSU1 and SSU3)

494

and IPMDH1; a schematic diagram showing the assembly of all genes involved in

21



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

495

glucoraphanin biosynthesis in vivo; RT-PCR analyses of the gene expression in the

496

glucoraphanin biosynthesis pathway in E. coli; and the sequences of all genes used in

497

this study supplied as Supporting Information.

498

499 500 501

Author Contributions B.Y. and Y. L. designed the research. H.Y. and F.L. performed the experiments. H.Y., Y. L. and B.Y. wrote the article.

502 503 504

Notes The authors declare no competing financial interest.

505 506 507 508

Acknowledgements The work was partially supported by a grant from the Key International Cooperation Project of Chinese Academy of Sciences (155112KYSB20150024).

509 510

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511

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Mortensen, U. H., and Halkier, B. A. (2012) Microbial production of

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610

(32) He, Y., Chen, B., Pang, Q., Strul, J. M., Chen, S. (2010) Functional specification of

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29



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Page 30 of 42

Table 1 Primers used in protein expression in this study. Primers

Sequences (5’-3’)*

Function

BCAT3-F

CCCAAGCTTATGAATGCCGTGCTGAGCAATAGCAGC

Expression of truncated

BCAT3-R

CCGCTCGAGTGTTAACACGGTATACAG

BCAT3

GSL-ELONGF

CGGAATTCATGCCGCCGCAGAAGATCGAAATTGCCC


GSL-ELONGR

CCGCTCGAGCACCACGCTGCTGATCTG

GSL-ELONG

IPMI-LSU1-F

CGGAATTCATGACAATGACGGAGAAGATTCTAG


IPMI-LSU1-R

ACCGCTACCGCTGCTGGTGCTACCCTGCAAGAACTCCCTTGGGTC

IPMI (LSU1&SSU3)

IPMI-SSU3-F

GGTAGCACCAGCAGCGGTAGCGGTATAACCAGAGAGACTTTCCAC

IPMI-SSU3-R

CCGCTCGAGTCAAGCAGAAGGAATCATGCCGGC

IPMDH-F

CGGAATTCATGGCTTCACCTGGGAAAAAACGG


IPMDH-R

CCGCTCGAGTTAAACAGTAGCTGGAACTTTGG

IPMDH1

79F1-F

ACGCGTCGACATGGCTCTGTTATTAGCAGTTTTTACCACCAGCCTGCCGTACCC

Expression of modified

79F1-R

ACCGCTACCGCTGCTGGTGCTACCCGGGCAAAACTTCGGATACAG

CYP79F1 fusing with

ATR2-F

GGTAGCACCAGCAGCGGTAGCGGTAGGAGATCCGGTTCTGGGAATTC

truncated ATR2

ATR2-R

CCGCTCGAGTTACCATACATCTCTAAGATATC

30


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1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48


83A1-F

ACGCGTCGACATGGCTCTGTTATTAGCAGTTTTTAGCCAGAAAAGCAAAACCAAAC

Expression of

83A1-R

ACCGCTACCGCTGCTGGTGCTACCTTTGCTAACTTTTTCCGGAACC

CYP83A1

EGT2-F

CCGGAATTCATGGTGGCAACCACCGTTGAAC

Expression of EGT2

EFT2-R

CCGCTCGAGTTATGCGCTTTCTTTGTAC

UGT74B1-F

CGCGGATCCATGGCCGAAACTACAACAAC

Expression of

UGT74B1-R

CCGCTCGAGTTAGTGTTTTTTGCCCAGAC

UGT74B1

ST5c-F

CGCGGATCCATGGAGAGCAAAAGCGAGAATG

Expression of ST5c

ST5c -R

CCGCTCGAGTTACGGGCTGCTTGCCAGAAAAC

FMO-F

CGAGCTCATGGCACCAACTCAAAACACAATC

Expression of

FMO-R

CCGCTCGAGTCATGATTCGAGGAAATAAGAAG

FMO-GSOX1

BMYR-F

CCGGAATTCATGGATTACAAATTTCCGAAAG

Expression of

BMYR-R

CCGCTCGAGTTACGGTTTGCCGGTGCTC

myrosinase

*

The restriction sites are undelined.

*

The linker sequences are bold.

31



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Page 32 of 42

Table 2 Primers used in the reconstitution of glucoraphanin biosynthetic pathway in E. coli. Sequences (5’-3’)*

Primers

Function

BCAT3-F

CATGCCATGGACATGAATGCCGTGCTGAGCAATAG

BCAT3-R

ATGTATATCCTCCTTATTATGTTAACACGGTATACAG

GSL-ELONGF

TAAGGAGGATATACATATGCCGCCGCAGAAGATCGAAATTG

GSL-ELONGR

CTAGCTAGCCACCACGCTGCTGATCTGCGGGCTC

IPMDH-F1

GGAATTGTGAGCGGATAACAATTCCTAAGGAGGATATACATATGGCTTCACCTGGGAAAA

Expression of BCAT3

Expression of GSL-ELONG

Expression of IPMDH1

AAC IPMDH-F2

CTAGCTAGCTAATACGACTCACTATAGGGGAATTGTGAGCGGATAACAATTCC

IPMDH-R

ACGCGTCGACTTAAACAGTAGCTGGAACTTTG

IPMI-LSU1-F

ACGCGTCGACTAAGGAGGATATACATATGACAATGACGGAGAAGATTCTAG

Expression of IPMI

IPMI-LSU1-R

ATGTATATCCTCCTTACTACTGCAAGAACTCCCTTGGGTC

(LSU1&SSU3)

IPMI-SSU3-F

TAAGGAGGATATACATATGATAACCAGAGAGACTTTCCAC

32


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IPMI-SSU3-R

CCGCTCGAGTCAAGCAGAAGGAATCATGCCGGC

79F1-F

CATGGCCGGCCATGGCTCTGTTATTAGCAGTTTTTACCAC

79F1-R

ATGTATATCCTCCTTATTACGGGCAAAACTTCGGATACAG

83A1-F

TAAGGAGGATATACATATGGCTCTGTTATTAGCAGTTTTTAG

83A1-R

ACATGCATGCTTATTTGCTAACTTTTTCCGGAACCAGTTTC

ATR2-F

ACATGCATGCTAAGGAGGATATACATATGGCTCTGTTATTAGCAGTTTTTAG

ATR2-R

CCTTAATTAATTACCATACATCTCTAAGATATCTTC

EGT2-F

CGAGCTCTAAGGAGGATATACATATGGTGGC

EFT2-R

TTTCGGCCATATGTATATCCTCCTTATTATGCGCTTTCTTTGTACTCGCC

UGT74B1-F

AAGCGCATAATAAGGAGGATATACATATGGCCGAAACTACAACAACAACC

UGT74B1-R

TCCCCCCGGGTTAGTGTTTTTTGCCCAGAC

ST5c-F

TCCCCCCGGGTAAGGAGGATATACATATGGAGAGCAAAAGCGAG

ST5c -R

TTGGTGCCATATGTATATCCTCCTTATTACGGGCTGCTTGCCAG

FMO3-F

CAGCCCGTAATAAGGAGGATATACATATGGCACCAACTCAAAAC

FMO3-R

ACGCGTCGACTCATGATTCGAGGAAATAAG

*

The restriction sites are undelined.

*

The RBS sequences are bold.

Expression of CYP79F1

Expression of CYP83A1

Expression of ATR2

Expression of EGT2

33


Expression of UGT74B1

Expression of ST5c

Expression of FMO3


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1

Figure Captions:

2

Fig. 1 Glucoraphanin biosynthesis pathway from methionine in plant.

3

The enzymes catalyze each step reaction are listed on the left column and full names for the enzymes

4

could be referred in the main text.

5 6

Fig. 2 Functional expression of modified cytochrome P450s in E. coli.

7

(a) Schematic diagram of constructing gene ensembles encoding the cytochrome CYP79F1 [8-540]

8

and CYP83A1 [20-501] with ATR2 [73-711], respectively. Numbers correspond to the amino acids

9

encoded by the first and last codons of the gene sequences. Synthetic sequences ε were used to

10

replace the deleted N-terminal domain of the native P450s and the artificial linker λ was used for

11

fusion of P450s with ATR2.

12

(b) Functional expression of modified cytochrome P450 enzymes CYP79F1 and CYP83A1 fused

13

with reductase ATR2 in E. coli. The protein bands were indicated by the red arrows, respectively.

14 15

Fig. 3 Soluble expression of selected proteins involved in the core structure formation and secondary

16

modification pathway of glucoraphanin in E. coli.

17

The left is the diagram showing that EGT2 catalyzes the similar reaction with SUR1. The right is the

18

profile of showing the protein expression and the respective protein bands were indicated by the red

19

arrows.

20 21

Fig. 4 Detection of the reaction product from the core formation and side-chain modification pathway

22

in vitro.

23

(a) GC chromatogram of authentic standard sulforaphane.

24

(b) GC chromatogram of product from the enzyme mixture and the possible product was indicated

25

with an arrow. 34


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26

(c) Ion mass spectra of authentic standard sulforaphane and the structure was identified as 3-butenyl

27

isothiocyanate.

28

(d) Ion mass spectra of the product peak and the structure was identified accordingly.

29

(e) Thermal degradation of sulforaphane to 3-butenyl isothiocyanate caused by the high temperature

30

of the injection ports of GC and GC/MS.

31 32

Fig. 5 The biosynthesis pathway of glucoraphanin in Arabidopsis thaliana vs assembly in E. coli.

33

(a) Side-chain elongation of amino acid.

34

(b) Formation of core glucosinolate functional group.

35

(c) Secondary modification of side chain and breakdown of glucoraphane.

36 37

Fig. 6 Glucoraphanin production in E. coli by expression of the modified synthetic pathway.

38

(a) LC-MS/MS of authentic standard glucoraphanin. (b) Reaction product from the resting cell.

39

Multiple reaction monitoring (MRM) was used to monitor analyze parent ion production transitions.

40

MRMs were chosen as follow: glucoraphanin (m/z 436-96.8, 436-177.8, 436-259.0, 436-371.9).

35



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 1 Glucoraphanin biosynthesis pathway from methionine in plant 212x282mm (300 x 300 DPI)


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Figure 2 Functional expression of modified cytochrome P450s in E. coli 80x61mm (300 x 300 DPI)



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 3 Soluble expression of selected proteins involved in the core structure formation and secondary modification pathway of glucoraphanin in E. coli 140x69mm (300 x 300 DPI)


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Figure 4 Detection of the reaction product from the core formation and side-chain modification pathway in vitro 140x108mm (300 x 300 DPI)



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 5 The biosynthesis pathway of glucoraphanin in Arabidopsis thaliana vs assembly in E. coli 80x74mm (300 x 300 DPI)


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Figure 6 Glucoraphanin production in E. coli by expression of the modified synthetic pathway 80x109mm (300 x 300 DPI)



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Abstract Graphic 69x39mm (300 x 300 DPI)


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Reconstructing Biosynthetic Pathway of the Plant ... - ACS Publications

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