- Open Access
Biosynthesis of dendroketose from different carbon sources using in vitro and in vivo metabolic engineering strategies
© The Author(s) 2018
- Received: 19 June 2018
- Accepted: 15 October 2018
- Published: 25 October 2018
Asymmetric aldol-type C–C bond formation with ketones used as electrophilic receptor remains a challenging reaction for aldolases as biocatalysts. To date, only one kind of dihydroxyacetone phosphate (DHAP)-dependent aldolases has been discovered and applied to synthesize branched-chain sugars directly using DHAP and dihydroxyacetone (DHA) as substrate. However, the unstable and high-cost properties of DHAP limit large-scale application. Therefore, biosynthesis of branched-chain sugar from low-cost and abundant carbon sources is essential.
The detailed catalytic property of l-rhamnulose-1-phosphate aldolase (RhaD) and l-fuculose-1-phosphate aldolase (FucA) from Escherichia coli in catalyzing the aldol reactions with DHA as electrophilic receptors was characterized. Furthermore, we calculated the Bürgi–Dunitz trajectory using molecular dynamics simulations, thereby revealing the original sources of the catalytic efficiency of RhaD and FucA. A multi-enzyme reaction system composed of formolase, DHA kinase, RhaD, fructose-1-phosphatase, and polyphosphate kinase was constructed to in vitro produce dendroketose, a branched-chain sugar, from one-carbon formaldehyde. The conversion rate reached 86% through employing a one-pot, two-stage reaction process. Moreover, we constructed two artificial pathways in Corynebacterium glutamicum to obtain this product in vivo starting from glucose or glycerol. Fermentation with glycerol as feedstock produced 6.4 g/L dendroketose with a yield of 0.45 mol/mol glycerol, representing 90% of the maximum theoretical value. Additionally, the dendroketose production reached 36.3 g/L with a yield of 0.46 mol/mol glucose when glucose served as the sole carbon resource.
The detailed enzyme kinetics data of the two DHAP-dependent aldolases with DHA as electrophilic receptors were presented in this study. In addition, insights into this catalytic property were given via in silico simulations. Moreover, the cost-effective synthesis of dendroketose starting from one-, three-, and six-carbon resources was achieved through in vivo and in vitro metabolic engineering strategies. This rare branched-chain ketohexose may serve as precursor to prepare 4-hydroxymethylfurfural and branched-chain alkanes using chemical method.
- Aldol reactions
- Branched-chain sugar
- Metabolic engineering
Directed aldol reaction is one of the most powerful carbon–carbon bond-forming procedures in synthetic organic chemistry and enables the concomitant creation of functionalized stereogenic centers and construction of chiral complex polyhydroxylated molecules [1, 2]. Catalytic asymmetric addition of carbon nucleophiles (donor) to ketones (acceptor) is a fundamental approach to construct new tetrasubstituted stereogenic carbon centers. This reaction is synthetically efficient to synthesize chiral tertiary alcohols, which are important building blocks of naturally occurring and artificial biologically active molecules . Aldol addition with ketones as electrophilic receptors is extremely challenging compared with the catalytic enantioselective aldol reaction to aldehydes. Few successful examples have been shown in catalytic aldol reaction to ketones, and they rely on metal catalysts or highly reactive trichlorosilyl enolate of methyl acetate [4–6]. However, all these reactions have a narrow substrate scope and depend on activated ketone acceptors or chiral auxiliaries.
Biocatalyzed aldol additions are attractive because this type of reaction occurs under mild conditions. As such, aldolases are particularly compatible as catalysts in the production of chiral compounds due to high selectivity and catalytic efficiency [7, 8]. A number of aldolases for catalyzing enantioselective aldol additions taking aldehydes as acceptors have been developed . However, to date, only two aldolases that can use ketones as acceptors have been reported. One is pyruvate-dependent aldolase from Pseudomonas taetrolens, which exhibits the catalytic ability in aldol addition of pyruvate to a ketone acceptor indole-pyruvic acid and has been used in the stereoselective synthesis of a precursor of monatin [10, 11]. The other one is l-rhamnulose-1-phosphate aldolase (RhaD) from Bacteroides thetaiotaomicron. This enzyme catalyzes the aldol reaction between DHAP and several ketones (hydroxyacetone, 1-hydroxybutanone, hydroxypyruvate and l-erythrulose) and gave four branched-chain sugars by coupling an acid phosphatase . The development of several kinds of aldolases that can tolerate ketones as electrophilic receptors is necessary in the asymmetric catalysis field.
Branched-chain sugars, e.g., dendroketose, which belong to a class of rare sugars, are monosaccharides, which rarely exist in nature . Dendroketose was obtained from the polymerization of two molecules of dihydroxyacetone [14, 15]. Typically, dendroketose that contains a tertiary alcohol moiety can be chemically dehydrated to furfural derivatives 4-hydroxymethylfurfural (4-HMF) which showed broad application prospects in preparing fine chemicals . Such conversion is similar to that of fructose to 5-hydroxymethylfurfural (5-HMF) . The synthesis of 2,5-dimethylfuran (DMF) from 5-HMF is a highly attractive route to a renewable fuel . The feasibility of producing 2,4-dimethylfuran (2,4-DMF) or C9–C15 branched-chain alkanes as liquid transportation fuels from 4-HMF has also been demonstrated . Therefore, developing methods that can synthesize branched-chain sugar is meaningful. The asymmetric aldol addition reactions via DHAP-dependent aldolases are particularly striking strategies in preparation of innovative branched-chain sugars due to the direct and rapid creation of molecular complexity in benign environments . DHAP-dependent aldolases exhibit a strict specificity for the donor DHAP . However, compound DHAP is unstable  and currently very expensive. These conditions limit their large-scale application.
In this work, we investigated the catalytic properties of two DHAP-dependent aldolases with ketones as electrophilic receptors. Moreover, we calculated the Bürgi–Dunitz trajectory using molecular dynamics (MD) simulations to reveal the original sources of the catalytic efficiency. Finally, an in vitro multi-enzyme system was designed, and two in vivo artificial pathways were constructed in Corynebacterium glutamicum to synthesize dendroketose from one-, three-, and six-carbon resources.
Aldol reactions to DHA catalyzed by DHAP-dependent aldolases
Steady-state kinetic parameters of aldol reactions to DHA catalyzed by RhaD and FucA
103 × kcat/KM (s−1 mM−1)
4.3 ± 0.5
40.8 ± 5.8
20.5 ± 2.5
500 ± 64
0.86 ± 0.24
23.4 ± 9.69
4.1 ± 0.52
180 ± 40
1.5 ± 0.3
54.1 ± 3.6
5.0 ± 0.53
92 ± 13
0.32 ± 0.13
309.8 ± 45.2
0.45 ± 0.13
1.4 ± 0.5
Molecular dynamic simulations provide insights into the catalytic properties of RhaD and FucA
To gain insights into the catalytic properties of RhaD and FucA from E. coli, we determined the apparent steady-state kinetic parameters in the reactions of the nucleophile DHAP and acceptor DHA. Substrate l-glyceraldehyde (l-GAL) was evaluated as the control to compare the catalytic properties when using aldehyde and ketone as acceptors. Despite the lower KM value of RhaD to DHA (23.4 mM) than that of l-GAL (40.8 mM), the kcat/KM value of RhaD to DHA was threefold lower than that of l-GAL (Table 1). These results indicated that DHA binds to RhaD with higher affinity but with lower catalytic activity than l-GAL. In the case of FucA, the kcat/KM value of FucA to DHA was 65-fold lower than that of FucA to l-GAL. This result indicated that the catalytic activity of FucA toward DHA was significantly lower than that to l-GAL. In terms of kcat/KM, clearly, the value of RhaD to DHA was 350-fold higher than that of FucA to DHA. This finding suggested that RhaD is much more efficient in catalyzing the aldol reaction between DHAP and DHA than FucA. According to the KM value of FucA in Table 1, the failure of FruA and TagA in catalyzing the aldol addition to DHA was probably due to very low affinity and catalytic efficiency to this substrate.
Construction of in vitro multi-enzyme system to produce dendroketose from formaldehyde
The key information of enzymes used in multi-enzyme system
Specific activities (U/mg)
l-Rhamnulose-1-phosphate aldolase (RhaD)
DHA kinase (DhaK)
Polyphosphate kinase (PPK)
When 40 mM FALD was used, the system produced 12.1 mM DHA with a conversion rate of 91%. This value was calculated with the ratio of DHA formation amount to initial concentration of FLAD. The GA has not been detected in this system. This result was identical to the previous study for which DHA was the primary product at high concentrations of FALD (> 10 mM) . In the second stage, the individually purified enzymes DhaK, RhaD, YqaB, PPK, 10 mM polyphosphate, 0.5 mM ATP and 5 mM MgSO4 were added into the reaction system. After reaction for another 24 h, this system produced 5.7 mM (1.03 g/L) dendroketose with a conversion of 86%, which was calculated with the ratio of dendroketose formation amount to initial concentration of FLAD (Fig. 5b). We also detected a small amount of d-erythrulose (~ 0.5 mM) which was derived from the aldol reaction between DHAP and FALD. In this multi-enzyme system, the enzyme activity of FLS to FALD was low for DHA production. Combination of mutational hot spots analytical method and site-saturated mutagenesis strategy has increased the catalytic efficiency of FLS to acetaldehyde for 72.9% . This protein engineering strategy showed the application potential in improving catalytic efficiency of FLS to FALD and further in increasing the production efficiency of dendroketose in the multi-enzyme system.
Pathway design and strain engineering to produce dendroketose from glycerol
Synthesis of dendroketose using different carbon sources
Glycerol and DHA
5.6 ± 0.4
0.68 ± 0.03
0.8 ± 0.1
0.05 ± 0.01
1.2 ± 0.2
0.46 ± 0.02
6.4 ± 0.4
0.45 ± 0.02
36.3 ± 1.3
0.46 ± 0.02
8.2 ± 0.6
0.12 ± 0.02
Metabolic engineering of dendroketose production from glucose
Strains and plasmids used in this study
Strains and plasmids
Genotype and properties
Source or references
E. coli DH5α
supE44 ΔlacU169 (φ80 lacZΔM15) hsdR17 recA1 endA1 gyrA96 thi-1 relA1
E. coli BL21 (DE3)
F-ompT hsdSB (Rb-mB-) gal (λ c I 857 ind1 Sam7 nin5 lacUV5 T7gene1) dcm (DE3)
Gene tpi knock-out in C. glutamicum 13032
Wide-type containing plasmid pXRTYH
Strain SY6 containing plasmid pXRTYH
Strain SY6 containing plasmid pXFucTYH
Wide-type containing plasmid pXRTY and pEFDK
Strain SY6 containing plasmid pXRTY and pEFDK
Expression vector, ApR
pET21a(+) derivative carrying gene of rhaD
pET21a(+) derivative carrying gene of fucA
pET21a(+) derivative carrying gene of FLS
pET21a(+) derivative carrying gene of yqaB
pET21a(+) derivative carrying gene of DhaK
pXMJ19 derivative carrying gene of rhaD and yqaB
pECXK99E derivative carrying gene of glpF, dhaD and dhaK
pXRTY derivative carrying gene of hdpA
pXFucTY derivative carrying gene of hdpA
In summary, the detailed catalytic property of RhaD and FucA from E. coli in direct aldol addition using DHA as electrophile acceptor was characterized. The Bürgi–Dunitz trajectory calculated by MD simulations has been applied to reveal the catalytic efficiency difference of those two aldolases. Furthermore, we provided a green and environment-friendly biocatalytic approach to synthesize a rare branched-chain sugar dendroketose directly from FALD with high conversion rate. The synthesis of dendroketose from renewable feedstock glucose or glycerol using an engineered strain was also achieved with high titre and yield. Those advanced routes developed in this study presented low-cost way in producing dendroketose. Chemical dehydration of dendroketose will obtain 4-HMF, which serves as a potential platform molecule in preparing certain biofuels and fine chemicals.
Bacterial strains, plasmids and materials
Compounds DHAP, DHA, l-GAL, AP from potato, isopropyl-β-d-thiogalactopyranoside (IPTG), polyphosphate, ATP, l-fructose, l-tagatose, glycerol, glucose, and antibiotics were purchased from Sigma-Aldrich. All restriction enzymes and DNA ligase were purchased from Novagen (Darmstadt, Germany). Ni–NTA affinity chromatography column was purchased from QIAGEN. The yeast extract and tryptone were purchased from OXOID LID, and brain heart infusion (BHI) was purchased from Becton, Dickinson and Company. All bacterial strains and plasmids are listed in Table 4.
Vectors and strains construction
The genes fucA from E. coli and dhaK from Citrobacter freundii were amplified from genome and cloned into pET-21a(+) to obtain pET21-FucA and pET21-DhaK, respectively. The plasmid pET21-PPK containing the gene of ppk from Rhodobacter sphaeroides was kindly provided by Professor Chun You in our institute. For the construction of plasmid pXRTYH and pXFucTYH, gene hdpA was amplified from C. glutamicum 13032 genome. The amplified fragments were ligated into plasmid previous constructed plasmids pXRTY  and pXFucTY  at the SmaI and SacI sites to obtain pXRTYH and pXFucTYH, respectively. The constructed plasmids were then electroporated into the recombinant strain SY6, in which the gene tpi has been eliminated, to generate strains SY6(pXRTYH) and SY6(pXFucTYH). Plasmid pEFDK containing genes glpF, dhaD and dhaK  and pXRTY were co-transformed into SY6 strain to obtain SY6(pXRTY/pEFDK).
Recombinant proteins expression and purification
Escherichia coli BL21(DE3) strains harboring expression plasmids were cultured at 37 °C in 1 L LB medium containing 100 mg/L ampicillin to an optical density OD600 of 0.6. 0.5 mM IPTG was added into the culture to induce protein expression and the temperature was adjusted to 16 °C to avoid inclusion body formation. After incubation for an additional 20 h, cells were harvested, washed twice and suspended in 50 mM triethanolamine (TEA) (pH 7.5) buffer. The suspension cells were then lysed by sonication and centrifuged at 14,000×g and 4 °C for 10 min. Clear supernatant was collected and loaded onto an Ni2+-NTA-agarose column pre-equilibrated with binding buffer (50 mM TEA buffer, 300 mM NaCl, 20 mM imidazole, pH 7.5). The retained proteins were recovered with elution buffer (50 mM TEA buffer, 300 mM NaCl, 300 mM imidazole, pH 7.5). The eluted fraction containing purified protein was dialyzed to eliminate buffer, salt and imidazole. The purified enzymes were freeze dried using a vacuum pump and stored at − 20 °C.
Enzyme activity assay
The activity of PPK was assayed in a reaction mixture (200 μL) containing PPK (0.05 mg), 25 mM TEA buffer (pH 7.0), 25 mM DHA, 10 mM ADP, 10 mM polyphosphate and 5 mM MgCl2·6H2O. After the reaction at 30 °C for 30 min, the reaction was stopped by the addition of 10% H2SO4 (0.5 μL). Product was determined via high-performance liquid chromatography (HPLC). One unit of enzyme activity was defined as the enzyme amount catalyzing the consumption of 1 μmol DHA per min.
The activity of FLS was assayed in a reaction mixture (200 μL) containing FLS (2 mg), 25 mM TEA buffer (pH 7.0), 25 mM FALD, 1 mM MgSO4 and 0.1 mM TPP. After the reaction at 30 °C for 45 min, the reaction was stopped by the addition of 10% H2SO4 (0.5 μL). One unit of enzyme activity was defined as the enzyme amount catalyzing the formation of 1 μmol of total DHA and GA per min.
Steady-state kinetic parameters of RhaD and FucA to DHA and l-GAL
Reaction: aldol addition of DHAP to l-GAL. To a solution containing freshly neutralized DHAP (60 mM) and RhaD (0.05 mg powder) in 50 mM TEA buffer pH 7.5 at 25 °C, different amounts of l-GAL (0.2, 0.5, 2, 5, 10, 20, 40, 60 mM) were added. The final volume was 400 μL. Samples (40 μL) were withdrawn at different times (0, 2, 5, 10, 20, 30 min) and the reaction was stopped by the addition of 10% H2SO4 (0.5 μL). Samples were then analyzed by HPLC to measure the l-GAL consumption. One mmol of l-GAL consumed was equivalent to 1 mmol of l-fructose-1-phosphate formed.
Reaction: aldol addition of DHAP to DHA. To a solution containing freshly neutralized DHAP (60 mM) and RhaD (0.05 mg powder) in 50 mM TEA buffer pH 7.5 at 25 °C, different amounts of DHA (2, 5, 10, 20, 40, 60, 80 mM) were added. The final volume was 400 μL. Samples (40 μL) were withdrawn at different times (0, 2, 5, 10, 20, 30 min) and the reaction was stopped by the addition of 10% H2SO4 (0.5 μL). Samples were then analyzed by HPLC to measure the DHA consumption. One mmol of DHA consumed was equivalent to 1 mmol of adduct formed.
Reaction: aldol addition of DHAP to l-GAL. To a solution containing freshly neutralized DHAP (60 mM) and FucA (0.1 mg powder) in 50 mM TEA buffer pH 7.5 at 25 °C, different amounts of l-GAL (0.5, 2, 5, 10, 20, 40, 60 mM) were added. The final volume was 400 μL. Samples (40 μL) were withdrawn at different times (0, 2, 5, 10, 20, 30 min) and the reaction was stopped by the addition of 10% H2SO4 (0.5 μL). Samples were then analyzed by HPLC to measure the l-GAL consumption. One millimole of l-GAL consumed was equivalent to 1 mmol of l-tagatose-1-phosphate formed.
Reaction: aldol addition of DHAP to DHA. To a solution containing freshly neutralized DHAP (60 mM) and FucA (0.1 mg powder) in 50 mM TEA buffer pH 7.5 at 25 °C, different amounts of DHA (2, 5, 10, 20, 40, 60, 80, 100, 200 mM) were added. The final volume was 400 μL. Samples (40 μL) were withdrawn at different times (10, 20, 40, 60, 90, 120, 180 min) and the reaction was stopped by the addition of 10% H2SO4 (0.5 μL). Samples were then analyzed by HPLC to measure the DHA consumption. One mmol of DHA consumed was equivalent to 1 mmol of adduct formed.
Aldol reactions with DHAP and DHA as substrates
The reaction mixture (1 mL) contained freshly neutralized 50 mM DHAP solution, 50 mM DHA, 50 mM TEA buffer (pH 7.5) and RhaD (1 mg) or FucA (2 mg). The reaction mixture was transferred to a 1.5-mL Eppendorf tube and shaken at 25 °C and 120 rpm for 24 h. Then, the pH of the mixture was adjusted to 4.5–5.5 using 10% H2SO4, and 2 U AP was supplemented. The dephosphorylation reaction was performed at 30 °C for another 24 h.
Models of the dimer structures of FucA complex and RhaD complex were generated as follows: the monomer structure of FucA is derived from the previous study (PDB code 4FUA). However, there is no available polymer structure of FucA by searching the PDB database. TM-align program  was used to search for FucA homologies; in the top 10 hits ranked by TM-score, 2OPI crystalized in polymers was used as template. In the case of RhaD, the X-ray structure (PDB code 1GT7) was directly used as the basis for dimer creation of RhaD. The coordinates of the donor DHAP and the acceptor DHA/L-GAL in constructed dimers were superimposed with those from the PDB codes 1OJR and 4FUA.
Based on the catalytic mechanism of class II Aldolase, a specific residue in the adjacent monomer (Tyr113′ in FucA, and Glu171′ in RhaD) plays a key role on the protonation of the carbonyl oxygen of ketone acceptors. Therefore, dimer models of enzyme–substrate complexes (dubbed as “FucA–DHAP–DHA”, “FucA–DHAP–GAL”, “RhaD–DHAP–DHA” and “RhaD–DHAP–GAL”) were built to reflect the catalytic mechanism.
During the simulations, a constant force of 10 kcal/mol between the nucleophile C-atom and the electrophile C-atom was constructed via the consideration of Van der Waals’ force. To estimate the stability of 100 ns trajectories of the four systems, root-mean-square deviation (RMSD) for all Cα atoms was analyzed and no significant structure difference was observed. Furthermore, root-mean-square fluctuation (RMSF) which could reflect the stability of individual residue of protein was also evaluated; most residues are stable except for the terminations between two monomers.
Molecular dynamics (MD) simulations
The initial structures used for MD simulation were obtained from modeling analysis. Each apo-protein was protonated at pH 7.5 using H++ webserver. The Amber ff14SB force field was employed for the protein in all the MD simulations . Na+ ions were added to neutralize the system, and the TIP3P water model was used to solvate each system, ensuring a solvent layer of at least 10 Å from any point on the protein surface. Charges and parameters for ligands were generated with the Antechamber module using the AM1-BCC charge model along with the amber GAFF force field. The force field of zinc ion and its neighboring atoms (cutoff was set as 2.8 Å) were parameterized using ‘MCPB.py’ modeling, using a hybrid bonded/restrained nonbonded model. As a result, the three histidine residues (H92, H94 and H155 in FucA, and H141, H143 and H212 in RhaD) were attached to zinc ion by coordinate bonds, whereas the two oxygen atoms of donor were attached to zinc ion by applying harmonic restraint (100 kcal/mol). After proper parameterizations and setup, the resulting system’s geometries were minimized (5000 steps for steepest conjugate and 5000 steps for conjugate gradient) to remove poor contacts and relax the system. The systems were then annealed from 0 to 300 K (≈ 27 °C) to mimic experimental temperature under the constant amount of substance (N), volume (V) and temperature (T) (NVT ensemble) for 50 ps. Subsequently, the systems were maintained for 25 ps of density equilibration under constant amount of substance (N), pressure (P) and temperature (T) (NPT ensemble) at constant temperature of 300 K and pressure of 1.0 atm using Langevin-thermostat (ntt = 3) with collision frequency of 2 ps−1 and pressure relaxation time of 1 ps. The heating and density equilibrations were carried out with a weak restraint of 20 kcal mol−1 Å−2 performed on all the residues. The systems were further equilibrated for 250 ps to get well settled pressure and temperature for conformational and chemical analyses. After proper minimizations and equilibrations, a productive MD run of 100 ns was performed for each system. During all MD simulations, the covalent bonds containing hydrogen were constrained using SHAKE algorithm , with a MD time step of 2 fs. The trajectory file was written every 1000 steps. All the above MD simulations were performed with GPU version of Amber 16 package. The generated trajectories (interval = 200, a total of 500 frames for each case) were used for the relative binding energy evaluation.
In vitro cascade reaction
To synthesize dendroketose in vitro, the reaction mixture (2 mL) containing 50 mM TEA (pH 7.5), 40 mM FALD, 0.1 mM TPP, FLS (1.5 U, 16 mg) was initially carried out at 25 °C and 120 rpm for 16 h. Then, DhaK (0.5 U, 0.025 mg), PPK (0.5 U, 0.03 mg), RhaD (0.5 U, 2 mg), YqaB (0.9 U, 1.5 mg), polyphosphate (20 mM), ATP (0.5 mM), and MgSO4 (10 mM) were added into the reaction system, and it performed for another 24 h. Samples (100 μL) were captured very two hours, treated with 10% H2SO4, centrifuged (22,000 rpm, 20 min) and analyzed by HPLC.
Shake flask scale cultivation
For precultivation of recombinant strain, a single clone was grown in 5 mL of BHI medium. After incubation for approximately 15 h, cells were inoculated into a 500-mL shake flask containing 100 mL BHI medium and cultivated at 25 °C in a rotatory shaker at 220 rpm. When the cell OD600 reached 0.8, 1 mM IPTG was added to induce enzyme expression. Subsequently, the cells were harvested by centrifugation (8000×g, 10 min, 4 °C) and were suspended in CGXII medium . Then, 50 mL cells were transferred into a 250-mL shake flask with an initial OD600 of approximately 30. When appropriate, 10 mg/L chloramphenicol and 25 mg/L kanamycin were added. The fermentation process was carried out at 30 °C and 200 rmp.
If glycerol and DHA were used as substrates, the concentration of glycerol and DHA was assigned to 220 mM and 110 mM, respectively. To produce dendroketose from glucose, 220 mM glucose was supplemented into the medium. For the fed-batch fermentation, 220 mM glucose was supplemented again into the medium after fermentation for 6 h. Samples were collected every 2 h and centrifuged at 14,000×g for 20 min. The resulting supernatants were analyzed by HPLC. The desired product was separated by a chromatographic column filled with Ca2+ ion exchange resin, identified by a refractive index detector and then collected by a fraction collector. The purified products were analyzed by NMR.
Cell density was determined by measuring the optical density at 600 nm (OD600) with a UV–Vis spectrophotometer (TU-1901, Persee, Beijing, China). Cell dry weight (CDW, g/L) of E. coli was calculated from OD600 values using the experimentally determined correlation factor of 0.25 g cells (dry weight [DW])/liter for an OD600 of 1. Protein concentrations were determined by the Bradford method using bovine serum albumin as a standard. HPLC system (Agilent 1100 series, Hewlett-Packard) equipped with a refractive index detector and fitted with chromatographic column (Bio-Rad Aminex HPX-87H column or Waters Sugar-Pak I column) was used to qualitative and quantitative analysis of substrates and products.
JY and YZ contributed equally to this work. They performed the experiments including enzyme screening, characterization and strain construction and drafted the manuscript. GQ performed the in silico simulations via molecular dynamics (MD). YZ, CT and CD carried out the experiments including product purification, MS and NMR analysis. YM and LD finished the fermentation experiments. ZS, YS and YM designed the constructs and revised the manuscript. All authors reviewed the results of the manuscript. All authors read and approved the final manuscript.
The authors declare that they have no competing interests.
Availability of data and materials
The dataset supporting the conclusions of this article is included within the article (and its Additional file).
Consent for publication
Ethics approval and consent to participate
This work was supported by the Natural Science Foundation of China (No. 21602245), Key Programs of the Chinese Academy of Sciences (No. ZDRW-ZS-2016-3), Youth Innovation Promotion Association of Chinese Academy of Sciences. Zhoutong Sun also thanks to the CAS Pioneer Hundred Talent Program (Type C) (reference number 2016-053) for initial start support.
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