CO2 fixation for malate synthesis energized by starch via in vitro metabolic engineering

Metabolic Engineering 55 (2019) 152–160

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CO2 fixation for malate synthesis energized by starch via in vitro metabolic engineering

T

Ting Shi, Shan Liu, Yi-Heng P. Job Zhang* Tianjin Institute of Industrial Biotechnology, Chinese Academy of Sciences, 32 West 7th Avenue, Tianjin Airport Economic Area, Tianjin, 300308, China

A R T I C LE I N FO

A B S T R A C T

Keywords: CO2 fixation Malate In vitro metabolic engineering Enzymatic cascade biocatalysis

Carbon dioxide (CO2) is an appealing carbon feedstock for the sustainable production of biocommodities. Here we designed three in vitro artificial enzymatic pathways featuring the ATP-excess, ATP-deficit, and ATP-balanced pathways for the biotransformation of starch and CO2 to malate. This ATP-balanced pathway without exogenous ATP donors can auto-regulate its carbon fluxes from glyceraldehyde 3-phosphate to 3-phosphoglycerate via either the ATP-generating pathway (a part of glycolysis) or no-ATP-generating pathway from a hyperthermophilic archaeon Thermococcus kodakarensis. The ATP-balanced pathway enabled to produce up to 52.4 mM malate with 95.3% of the theoretical yield, that is, 2 mol of malate synthesized from 1 mol of glucose of starch and 2 mol of CO2. This pathway also enabled to produce high-yield malate regardless of ATP/ADP ratios. Anaerobic reaction conditions and/or the addition of a reducing agent dithiothreitol were of importance for creating an anoxic environment for biocatalysis of enzyme cocktails and for mitigating the deactivation of enzymes and degradation of intermediates. This new pathway could provide a green route for direct conversion of CO2 to many building blocks, a promising alternative of petrochemical-based production of biocommodities.

1. Introduction Climate change has been linked to the accumulation of greenhouse gases (e.g., CO2) in the atmosphere caused by human activities, such as burning fossil fuels, land use changes, and deforestation (Claassens et al., 2016). The global average atmospheric CO2 increased to 405 ppm and global energy-related CO2 emissions reached a historic high level of 32.5 gigatonnes in 2017 (https://www.iea.org/topics/climatechange/). Hence, there is an urgent need to decrease the net emissions of CO2 to the atmosphere. The utilization of CO2 as a raw material in the synthesis of renewable fuels and chemicals offers a win-win strategy to both decrease the CO2 emissions and efficiently utilize this nearly free carbon resource (Venkata Mohan et al., 2016). Four major approaches for CO2 fixation and conversion reactions are chemical, photochemical, electrochemical and biological methods (Shi et al., 2015; Singh et al., 2018). Industrial chemical carbon utilization reactions include the synthesis of urea (dated back to 1870) and the Kolbe-Schmitt reaction for the production of salicylic acid (Lindsey and Jeskey, 1957) but their potential for CO2 utilization is limited by their small market sizes. It was urgent to find out more products that could be produced from CO2. Also, because CO2 has the highest oxidized state of carbon, a large external energy input is required for its chemical transformation (Glueck et al., 2010). Photochemical and electrochemical methods are *

in intensive investigation but they are far from technical maturity (Jhong et al., 2013; Kumar et al., 2012). Biological CO2 fixation is the most appealing strategy on a large scale due to its mild reaction conditions, great selectivity, high yield, and good scale-up potential. Plant photosynthesis is the predominant natural CO2 fixation process but it suffers from very low solar energy utilization efficiencies and slow volumetric productivity (i.e., requiring a lot of lands), as well as a large amount of water consumption (Zhang, 2013). A number of natural or engineered photoautotrophic microorganism featuring fast CO2 utilization rates and/or high energy utilization efficiencies have been investigated to produce biofuels and biochemicals, such as ethanol (Liang et al., 2018), 2,3-butanediol (Kanno et al., 2017), lipid (Ajjawi et al., 2017), ethylene (Xiong et al., 2015), lactic acid (Angermayr et al., 2012), and so on. Distinct from CO2 fixation energized by solar energy, biological CO2 fixation energized by pointed chemical energy sources exhibits much higher CO2 utilization rates and may have great industrial scale-up potentials (Lee et al., 2014). Anaerobic synthesis of succinic acid is one of the best examples of biological CO2 fixation powered by glucose. In theory, a yield of 1.71 mol of succinic acid can be synthesized from 1 mol glucose accompanied with 2 mol of CO2 when exogenous electrons are supplied (Liebal et al., 2018). However, practical yields of succinic acid were a little below this theoretic yield, such as 1.22 mol/mol glucose (Guettler

Corresponding author. E-mail address: [email protected] (Y.-H.P.J. Zhang).

https://doi.org/10.1016/j.ymben.2019.07.005 Received 15 May 2019; Received in revised form 11 July 2019; Accepted 11 July 2019 Available online 12 July 2019 1096-7176/ © 2019 Published by Elsevier Inc. on behalf of International Metabolic Engineering Society.

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balanced pathway can fix 2 mol CO2 by using 1 mol glucose unit from starch without exogenous addition of an ATP donor.

et al., 1999) and 1.5 mol/mol glucose (Zhu et al., 2014), the other carbon source was consumed for the synthesis of cell biomass and/or the formation of side-products. In vitro metabolic engineering (ivME) comprised of purified enzymes or cell lysates is an emerging biomanufacturing platform, beyond the constraints of living microorganisms, such as, net ATP generation used for their self-duplication and basic maintenance, the assimilation of small water soluble substrates (e.g., glucose) other than polymeric substrates (e.g., starch) across the cell membrane, and so on. Nearly theoretical yields of products can be implemented by ivME, for example, hydrogen production via advanced water splitting energized by numerous carbohydrates (Kim et al., 2018), 1,3-propandiol production from glycerol (Rieckenberg et al., 2014), lactate production from glycerol (Jaturapaktrarak et al., 2014), α-ketoglutarate production from glucuronate (Beer et al., 2017), myo-inositol from starch (Fujisawa et al., 2017; You et al., 2017), cellulose (Meng et al., 2018) and xylose (Cheng et al., 2019), and bioelectricity generation from glucose in biobatteries (Zhu and Zhang (2017)). Until now, a few studies have been reported about CO2 fixation and utilization by ivME, such as methanol (Singh et al., 2018) and glyoxylate (Schwander et al., 2016). However, efficient reduction of CO2 followed by carbon-carbon linkage for the synthesis of multi-carbon compounds are still one of essential questions for ivME. L-malate, a four-carbon dicarboxylic acid, is an essential metabolic intermediate in the tricarboxylic acid cycle and one of the major organic acids of apples and other fruits. It is one of the most important acidulants and flavor enhancers in beverage and food industries, and is also widely used in metal cleaning, textile finishing, cosmetic, pharmaceuticals, and synthesis of various fine chemicals. The U.S. Department of Energy has identified L-malate as one of the top 12 building block chemicals, which can potentially be produced from renewable biomass on a large industrial scale (Werpy and Petersen (2004)). Currently, the global demand of malate is estimated to be approximately 200,000 tons per year, while the annual production of malate is 40,000 tones (Chi et al., 2016). At its current selling prices of approximately ~10 RMB/kg, four-time that of starch (i.e., ~2.5 RMB/ kg), it could be profitable to make L-malate from starch and CO2 if the highly-efficiently biotransformation was available. L-malate was extracted originally from apple juice but this method suffers from high production costs (Tsao et al., 1999). Currently, it can be produced by three methods. First, it can be synthesized via the hydration of either maleate or fumarate at high pressure and temperature (Goldberg et al., 2006) but it yields a racemic mixture of D- and L-isomers, which are not suitable for food and pharmaceutical applications. Second, it can be synthesized via enzymatic hydration of fumarate by immobilized fumarase or whole-cell (Giorno et al., 2001; Presecki and Vasic-Racki, 2005), but fumarate is made from non-renewable petrochemical feedstock, resulting in net CO2 emissions. Third, microbial fermentation of malate is an appealing alternative (Chi et al., 2016). Its aerobic fermentation has a theoretical low yield of malate (i.e., 1 mol malate per mole glucose), accompanied with CO2 as a by-product. Anaerobic fermentation of malate might have a high theoretical yield (i.e., 2 mol malate per mole glucose), but this pathway, unlike glucose-to-lactate pathway that produces two molecules of lactate per molecule of glucose plus two molecules of ATP, does not generate net ATP to maintain basic cellular metabolism and self-duplication (Shaw et al., 2016; Zhang et al., 2011). To address zero-ATP production in anaerobic malate biotransformation, a two-stage bioprocess was developed: (1) aerobic fermentation produced enough ATP for the synthesis of cell biomass; and (2) anaerobic whole-cell biotransformation converted glucose to malate without ATP consumption (Zhang et al., 2011). However, the practical yield (1.42 mol malate/mol glucose) was still far below the theoretical yield (Liu et al., 2017; Zhang et al., 2011). In this study, we designed the three in vitro artificial enzymatic pathways to produce malate from starch and CO2 featuring ATP-excess, ATP-deficit, and their combination – called ATP-balanced. This ATP-

2. Materials and methods 2.1. Chemicals All chemicals used were of analytical grade or higher quality and purchased from Sigma-Aldrich (St. Louis, MO, USA), Aladdin (Shanghai, China) and Sinopharm (Beijing, China) unless specified. PrimeSTAR Max DNA Polymerase (Takara, Dalian, China) was used for PCR. The Luria-Bertani (LB) medium was used for Escherichia coli cell growth and recombinant protein expression. The final concentration of antibiotic for E. coli was 100 mg/L ampicillin. Maltodextrin (dextrose equivalent (DE) = 4.0–7.0) from Sigma-Aldrich has a number-average degree of polymerization (DP) of 23.8, measured by the method described elsewhere (You et al., 2017). 2.2. Construction of plasmids The strains and plasmids used in this study are listed in Table S1. The primers (Table S2) used for PCR amplification were synthesized by GENEWIZ (Beijing, China). Plasmid pET20b-StIA encoding His-tagged isoamylase (IA, E.C. 3.2.1.68) from archaeon Sulfolobus tokodaii was described previously (Cheng et al., 2015). Three plasmids pET20bTmαGP encoding alpha-glucan phosphorylase (αGP, EC 2.4.1.1) from Thermotoga maritima MSB8, pET20b-TkPGM encoding phosphoglucomutase (PGM, E.C. 5.4.2.2) from Thermococcus kodakarensis KOD1, pET20b-TtcPGI encoding glucose 6-phosphate isomerase (PGI, E.C. 5.3.1.9) from Thermus thermophilus HB27 were described previously (Wang et al., 2017). Plasmid pET20b-Ti4GT encoding 4-glucanotransferase (4GT, EC 2.4.1.25) from Thermococcus litoralis was described previously (You et al., 2017). Plasmid pET28a-TfuPPGK encoding the engineered polyphosphate glucokinase (PPGK, E.C. 2.7.1.63) gene from Thermobifda fusca was described previously (Zhou et al., 2018). The other expression plasmids were constructed from the pET20b(+) plasmid backbone using Simple Cloning (You et al., 2012). The genes encoding fructose-bisphosphate aldolase (ALD), triosephosphate isomerase (TIM), phosphoglycerate kinase (PGK), enolase (ENO) and phosphoenolpyruvate carboxylase (PEPC) were amplified by PCR from the chromosomal DNA of T. thermophilus HB27 to construct respective expression plasmids. The genes encoding glyceraldehyde-3-phosphate dehydrogenase (GADPH) and malate dehydrogenase (MDH) were amplified by PCR from the chromosomal DNA of T. maritima MSB8 and Archaeoglobus fulgidus to construct expression plasmids, respectively. The genes encoding 6-phosphofructokinase (PFK) from T. thermophilus HB8, non-phosphorylating glyceraldehyde-3-phosphate dehydrogenase (GAPN) from T. kodakarensis KOD1 and cofactor-independent phosphoglycerate mutase (PGAM) from Pyrococcus horikoshii OT3 were amplified by PCR from the vector pGETS-CGP which was gifted from professor K. Honda of Osaka University (Japan) (Ninh et al., 2015) to construct respective expression plasmids. 2.3. Overexpression and purification of recombinant proteins E. coli BL21(DE3) was used for overexpression of recombinant proteins. E. coli BL21(DE3) harboring the expression vector was cultured in the LB medium supplemented with 100 mg/L of ampicillin with a rotary shaking rate of 250 rpm at 37 °C. Protein expression was induced at an absorbance of ~1.0 at 600 nm by adding 0.1 mM isopropyl-β-D-thiogalactopyranoside (IPTG) and the cultivation temperature was decreased to 18 °C for another 16–20 h cultivation. Cells were harvested by centrifugation at 4 °C, washed twice with 50 mM 4-(2hydroxyethyl)-1-piperazineethane sulfonic acid (HEPES) buffer (pH 7.5), and re-suspended in 50 mM HEPES buffer (pH 7.5). Re-suspended cells were lysed by using sonification in an ice bath (Fisher Scientific 153

154 Pyrococcus horikoshii OT3 T. thermophilus HB27 T. thermophilus HB27

PGI

PFK ALD TIM GAPDH

PGK GAPN

PGAM

ENO PEPC

MDH

glucose 6-phosphate isomerase

6-phosphofructokinase fructose-bisphosphate aldolase triosephosphate isomerase glyceraldehyde-3-phosphate dehydrogenase phosphoglycerate kinase non-phosphorylating glyceraldehyde-3phosphate dehydrogenase cofactor-independent phosphoglycerate mutase enolase phosphoenolpyruvate carboxylase

Malate dehydrogenase

Estimated at 50 °C.

T. thermophilus HB27 T. kodakarensis KOD1

PGM

phosphoglucomutase

a

Thermotoga maritima MSB8 Thermococcus kodakarensis KOD1 Thermus thermophilus HB27 T. thermophilus HB8 T. thermophilus HB27 T. thermophilus HB27 T. maritima MSB8

αGP

alpha-glucan phosphorylase

Archaeoglobus fulgidus

Source

Abb.

Enzyme

Table 1 Hyperthermophilic enzymes used in this study.

1.1.1.37

4.2.1.11 4.1.1.31

5.4.2.12

2.7.2.3 1.2.1.90

2.7.1.11 4.1.2.13 5.3.1.1 1.2.1.12

5.3.1.9

5.4.2.2

2.4.1.1

EC

100 6 30

oxaloacetate + NADH = malate + NAD+

35

130 2

5 18 435 22

49

6

4

Sp. Act.(U mg−1)a

2-phospho-D-glycerate = phosphoenolpyruvate + H2O Phosphoenolpyruvate + CO2 + H2O =Orthophosphate + oxaloacetate

2-phospho-D-glycerate = 3-phospho-D-glycerate

ADP + 3-phospho-D-glyceroyl phosphate = ATP + 3-phospho-D-glycerate D-Glyceraldehyde 3-phosphate + NAD+ + H2O = 3-phospho-D-glycerate + NADH

ATP + D-fructose 6-phosphate = ADP + D-fructose 1,6-bisphosphate D-fructose 1,6-bisphosphate = glycerone phosphate + D-glyceraldehyde 3-phosphate D-Glyceraldehyde 3-phosphate = glycerone phosphate D-glyceraldehyde 3-phosphate + phosphate + NAD+ = 3-phospho-D-glyceroyl phosphate + NADH + H+

D-Glucose 6-phosphate = D-fructose 6-phosphate

D-Glucose 1-phosphate = D-Glucose 6-phosphate

(1,4-alpha-D-glucosyl)n + phosphate=(1,4-alpha-D-glucosyl)n-1 + alpha-D-glucose 1-phosphate

Reaction

Ninh et al. (2015) Nakamura et al. (1996) Langelandsvik et al. (1997)

Nojima et al. (1979) Matsubara et al. (2011) Ninh et al. (2015)

Ninh et al. (2015) Myung et al. (2014) Myung et al. (2014) Wrba et al. (1990)

Wang et al. (2017)

Wang et al. (2017)

Wang et al. (2017)

Ref.

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1 mM thiamine pyrophosphate (TPP), 2 mM ATP, 2 mM NAD+, 10 mM MgCl2, 0.5 mM MnCl2 and 100 mM NaHCO3. The three reactions were performed at 50 °C for 24 h. An aliquot (65 μL) of the reaction sample was withdrawn and then mixed with 35 μL of 1.88 M HClO4 for stopping the reaction. The pH value of the reaction solution was adjusted to neutral with 13 μL of 5 M KOH. The concentrations of malate in supernatants were examined by HPLC. Reaction buffers with different pH values, such as Citrate (pH 5.0 to 6.5), MOPS (pH 6.5 to 8.0), HEPES (pH 7.0 to 8.5), Tris-HCl (pH 7.0 to 9.0), were tested at 50 °C for 6 h. The HEPES concentrations were tested at 50, 100, 200 and 400 mM at 50 °C for 6 h to find out the optimal buffer concentration. The reaction temperatures of 40, 50, 60, and 70 °C was carried out in 100 mM HEPES buffer (pH 7.5) to find out the optimal temperature. The reaction solutions with different redox states of aerobic, aerobic supplemented with 2 mM dithiothreitol (DTT), anaerobic, and anaerobic supplemented with 2 mM DTT were tested at 50 °C to find out the optimal reaction condition. Meanwhile, a redox indicator resazurin, which turned a trace pink to clear color under anoxic condition, was added into the reaction solution to determine whether the molecular oxygen was removed completely. Strictly anaerobic conditions for one-pot malate synthesis were conducted in the Hungate culture tubes. The aqueous mixtures were vacuumed and flushed by ultrapure nitrogen for three times, as described elsewhere (Song et al., 2019). The residual dissolved oxygen in the aqueous solution was removed in an anaerobic chamber. To find the rate-limiting enzymes and optimize enzyme loadings, we increased a single enzyme concentration by 2-fold or 5-fold while keeping the concentrations of the other enzymes constant. The malate concentrations were measured after 6 h reactions. Furthermore, 1, 2, 5, 7.5, 10, 12.5, and 15 U/mL enzyme cocktails were used for malate production. For the complete starch utilization to malate production, 27.5 mM glucose equivalent maltodextrin was treated with IA, 4GT and PPGK were added sequentially and 5 mM polyphosphate was added with the addition of PPGK.

Sonic Dismembrator Model 500; 3 s pulse on and 6 s off, total 300 s at 50% amplitude). After centrifugation, the supernatant was purified by heat precipitation at 70 °C for 25 min. Protein concentration was measured by the Bio-Rad Bradford protein dye reagent method (Bio-Rad, Hercules, CA, USA) with bovine serum albumin as a reference. The purity of protein samples was examined by 12% SDS-PAGE. The purified enzymes were stored in 5% (w/v) glycerol at −80 °C. 2.4. Enzymatic activity assays All enzymes used in this study are summarized in Table 1. The specific activity of T. maritima αGP was measured in 20 mM PBS buffer (pH 7.5) containing 5 g/L maltodextrin, 5 mM MgCl2 as described elsewhere (Wang et al., 2017). The specific activity of T. kodakarensis PGM was measured in 100 mM HEPES buffer (pH 7.5) containing 5 mM G1P and 5 mM MgCl2, the product G6P was determined in 100 mM HEPES buffer (pH 7.5) containing 5 mM MgCl2, 5 mM NAD+ and 2 U of engineered thermostable Z. mobilis glucose 6-phosphate dehydrogenase (G6PDH) (Huang et al., 2018). The specific activity of T. thermophilus PGI activity was assayed in 100 mM HEPES buffer (pH 7.5) containing 5 mM MgCl2 with 5 mM fructose 6-phosphate (F6P) and the product G6P was determined as described in the PGM specific activity assay (Wang et al., 2017). The specific ATP-dependent activity of T. thermophilus PFK was measured in 100 mM HEPES buffer (pH 7.5) containing 5 mM F6P, 5 mM ATP, 5 mM MgCl2, 5 mM NAD+, 5 U ALD, 5 U TIM, 5 U GAPDH (Hansen et al., 2002). The specific activity of T. thermophilus TIM was measured in 100 mM HEPES buffer (pH 7.5) containing 5 mM MgCl2, 0.5 mM MnCl2, 5 mM glyceraldehyde 3-phosphate (G3P), 0.15 mM NADH, 1 U GAPDH (Myung et al., 2014). The specific activity of T. thermophilus ALD was measured in 100 mM HEPES buffer (pH 7.5) containing 5 mM MgCl2, 0.5 mM MnCl2, 2 mM G3P in the presence of TIM, FBP, and PGI (Myung et al., 2014). The specific activity of T. maritima GAPDH activity was assayed in 100 mM HEPES buffer (pH 7.5) containing 10 mM potassium dihydrogen arsenate, 3 mM NAD+ and 5.2 mM G3P (Wrba et al., 1990). The specific activity of PGK was measured in 100 mM HEPES buffer (pH 7.5) containing 1 mM EDTA, 1 mM ATP, 10 mM MgCl2, 10 mM 2-mercaptoethanol, 10 mM glycerate 3-phosphate, 0.2 mM NADH, and an excess amount of GAPDH (Nojima et al., 1979). The specific activity of T. kodakarensis GAPN was measured in 100 mM HEPES buffer (pH 7.5) containing 0.2 mM ATP, 5 mM MgCl2, 0.5 mM MnCl2, 5 mM NAD+, 5 mM G3P (Ninh et al., 2015). The specific activity of P. horikoshii PGAM was measured in 100 mM HEPES buffer (pH 7.5) containing 0.2 mM ADP, 5 mM MgCl2, 0.5 mM MnCl2, 0.2 mM NADH, 0.2 mM 3-phosphoglycerate (3-PG) and an excess amount of ENO, T. thermophilus pyruvate kinase, and T. thermophilus lactate dehydrogenase (Ninh et al., 2015). Similarly, the specific activity of ENO were spectrophotometrically evaluated in the mixture containing the substrate of 0.2 mM 2-phosphoglycerate (2-PG) instead of 3-PG. The specific activity of T. thermophilus PEPC was measured in 100 mM HEPES buffer (pH 7.5) containing 0.4 mM PEP, 10 mM KHCO3, 5 mM MgCl2, 0.4 mM NADH, 0.3 mM acetyl-CoA, and 5 U MDH (Nakamura et al., 1996). The specific activity of A. fulgidus MDH was measured in 100 mM HEPES buffer (pH 7.5) containing 0.4 mM NADH and 0.4 mM oxaloacetate (Langelandsvik et al., 1997).

2.6. Calculation of malate yields One mole of glucose 6-phosphate (G6P) generated from starch can fix 2 mol of CO2 and generates 2 mol of malate (Fig. 1C). In theory, 27.5 mM glucose equivalent IA-treated maltodextrin can produce 27.5 mM G6P and finally catalyzes by the enzyme cascades to produce 55 mM malate with 55 mM CO2 fixation. In practice, the amount of malate generated depends on the G6P amount made from maltodextrin. The theoretical maximum yield was 2 mol malate per mole glucose. 2.7. Other assays Malate was quantified by HPLC equipped with a Bio-Rad 87H column and a differential refraction detector. The column at 35 °C was eluted with a mobile phase of 5 mM H2SO4 at a flow rate of 0.5 ml/min. The concentration of maltodextrin was measured by the starch assay kit (Novozymes, Copenhagen, Denmark). The purities of heat-treated recombinant proteins were checked by SDS-PAGE and analyzed by using a densitometry analysis of the Image Lab software (Bio-Rad, Hercules, CA, USA).

2.5. One-pot biosynthesis of malate

3. Results

The 13-enzyme cocktails at their different ratios were reconstituted followed by ultra-filtration against 100 mM HEPES buffer (pH 7.5) or other buffers (e.g., citrate, MOPS, Tris-HCl) in Millipore Amicon MWCO 3000 centrifugal filters. Ultra-filtration was carried out several times to ensure the residual glycerol level was below 0.1% glycerol, which came from the enzyme storage buffer. For the three proof-of-concept biotransformations, enzyme cocktails containing 1 U/mL of each enzyme were mixed with 200 mM MOPS buffer (pH 8.0) containing 27.5 mM glucose equivalent maltodextrin, 10 mM KH2PO4, 0.5 mM acetyl-CoA,

3.1. Design of in vitro artificial enzymatic pathways The malate synthesis pathway was designed to include three modules: Module 1, the production of glyceraldehyde 3-phosphate (G3P) from starch and phosphate catalyzed by six enzymes: alpha-glucan phosphorylase (αGP), phosphoglucomutase (PGM), 6-phosphate isomerase (PGI), ATP-dependent 6-phosphofructokinase (PFK), fructose155

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Fig. 1. Schematic of the synthetic biosystem for conversion starch and CO2 to malate via the ATP-excess pathway (A), ATP-deficit pathway (B) and ATPbalanced pathway (C). The enzymes used are alpha-glucan phosphorylase (αGP), phosphoglucomutase (PGM), 6-phosphate isomerase (PGI), ATP-dependent 6phosphofructokinase (PFK), fructose-bisphosphate aldolase (ALD), triosephosphate isomerase (TIM), glyceraldehyde-3-phosphate dehydrogenase (GAPDH), phosphoglycerate kinase (PGK), non-phosphorylating glyceraldehyde-3-phosphate dehydrogenase (GAPN), cofactor-independent phosphoglycerate mutase (PGAM), enolase (ENO), phosphoenolpyruvate carboxylase (PEPC), malate dehydrogenase (MDH). The metabolites are glucose 1-phosphate (G1P), glucose 6-phosphate (G6P), fructose 6-phosphate (F6P), fructose 1,6-diphosphate (F1,6-BP), glyceraldehyde 3-phosphate (G3P), 1,3-diphosphoglycerate (1,3-BPG), 3-phosphoglycerate (3-PG), 2-phosphoglycerate (2-PG), phosphoenolpyruvate (PEP), oxaloacetate (OAA), and inorganic phosphate (Pi).

Fig. 2. SDS-PAGE analysis of hyperthermophilic enzymes purified by heat treatment.

(Fig. 1B); and their combination (Fig. 1C). For Module 3 we chose the ATP-consuming PEPC that converts phosphoenolpyruvate (PEP) and CO2 to oxaloacetate (OAA) due to its negative Gibbs free energy (ΔG'° = −39.2 kJ/mol). When ATP-generating Module 2.1 was consolidated with Module 1 and 3, the entire pathway generated one ATP excess per glucose, called the ATP-excess pathway (Fig. 1A). When ATP-free Module 2.2 was consolidated with Modules 1 and 3, the entire pathway had one ATP deficit per glucose, called the ATP-deficit pathway (Fig. 1B). To ensure that in vitro ATP generation matched its consumption, we combined four Modules 1, 2.1, 2.2, and 3 to construct the ATP-balanced pathway (Fig. 1C). Modules 2.1 and 2.2 could self-adjust the carbon fluxes via

bisphosphate aldolase (ALD), and triosephosphate isomerase (TIM); Module 2, the production of 3-phosphoglycerate (3-PG) and NADH from G3P; and Module 3, the production of malate from 3-PG and CO2 catalyzed by four enzymes: cofactor-independent phosphoglycerate mutase (PGAM), enolase (ENO), phosphoenolpyruvate carboxylase (PEPC), and malate dehydrogenase (MDH). For Module 2, there are three design choices: Module 2.1 catalyzed by glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and phosphoglycerate kinase (PGK) can generate one ATP as a part of typical glycolysis (Fig. 1A); Module 2.2 catalyzed by non-phosphorylating glyceraldehyde-3-phosphate dehydrogenase (GAPN) that does not generate ATP in some hyperthermophilic archaea (e.g., T. kodakarensis KOD1) (Matsubara et al., 2011) 156

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oxygen-free conditions and/or the addition of reducing reagent were of importance for efficient biotransformation of ivME. To validate the robustness of the ATP-balanced pathway that could self-adjust carbon fluxes between Modules 2.1 and 2.2, the enzyme cocktails containing ATP only, ATP/ADP = 10:1; 5:1, 1:1; 1:5, 1:10, and ADP alone were investigated (Fig. 5A). The enzyme cocktails regardless of ATP levels produced comparative malate levels to validate the feasibility of the designed ATP regeneration and rebalancing between Modules 2.1 and 2.2. But the case of ADP only did not produce malate because the entire pathway cannot be operative at the beginning without initially-added ATP. When the ATP/ADP ratio decreased from 10:1 to 1:10, the total turn-over number increased from 2.11 to 20.5 (Fig. 5B). The increases in a single-enzyme loading by 2- and 5-fold did not significantly enhance malate titers (Fig. S4), suggesting there were no obvious rate-limiting steps. Furthermore, the malate titers were enhanced 1.8-fold and 2.4-fold when all enzyme loadings were increased by 2- and 5- folds, respectively (Fig. S4). When the total loadings of enzyme cocktails increased from 1 to 15 U/mL each (Fig. S5), the malate titer increased up to 27.1 mM. It meant that the malate yield was 0.986 mol/mol glucose. Cases of 15 U/mL and 12.5 U/mL did not show a significant difference in malate titers, indicating that this enzyme loading was sufficient.

Fig. 3. The proof-of-concept of malate production catalyzed by 1 U/ml enzyme cocktail.

either of modules without exogenous ATP donors. All the enzymes were selected from hyperthermophilic archaea and bacteria according to the Brenda database by considering their optimal reaction temperatures and specific activities. SDS-PAGE analysis shows the purity of recombinant enzymes produced in E. coli after heat precipitation (Fig. 2). The heat-treated enzymes were used for the following experiments.

3.3. Enhanced malate titers through the complete utilization of starch When non-treated (branched) maltodextrin was used, the malate titer was 27.0 mM (Fig. 6A) and its yield was approximately 50% of the theoretical yield (Fig. 6B). After isoamylase-treatment made linear amylodextrin, malate titer was increased to 38.6 mM with 70.2% of the theoretical yield (Fig. 6B). To increase amylodextrin utilization efficiency, 4-glucanotransferase (4GT) was added at hour 12 to generate more G1P from maltotriose and maltose, resulting in an increase of its yield from 70.2% to 90.4%. To utilize the residual glucose molecules released by 4GT, polyphosphate glucokinase (PPGK) along with polyphosphate was added at hour 24. After 48 h, the malate titer was increased to 52.4 mM (Fig. 6A) and the malate yield was increased to 1.91 mol/mol glucose, 95.3% of the theoretical maximum yield (i.e., 2 mol malate per mole glucose) (Fig. 6B). All these data suggested that all glucose units of maltodextrin can be used to energize CO2 fixation for malate synthesis.

3.2. Validation and optimization of one-pot biosynthesis The proof-of-concept experiments via the three pathways (Fig. 1) were conducted in 200 mM MOPS buffer (pH 8.0) containing 1 U/ml enzyme each. The ATP-excess, ATP-deficit, and ATP-balanced biosystems produced 0.60, 1.47, and 2.58 mM malate, respectively (Fig. 3 and Fig. S1). The ATP-balanced pathway lead to the highest malate titer, suggesting that efficient ATP balancing and recycling was of importance for the continuous running of in vitro synthetic pathways. Because reaction conditions could influence apparent activities of enzymes greatly (Ardao and Zeng, 2013), buffer type, pH, buffer concentration, reaction temperature, oxygen and reducing agent were optimized for the ATP-balanced system (Fig. S2). First, four kinds of buffers at different pH ranges, such as MOPS buffers (pH 6.5–8.0), HEPES buffers (pH 7.0–8.5), Tris-HCl buffers (pH 7.0–9.0), and citrate buffers (pH 6.0–6.5) were compared. The HEPES buffers (pH 7.0 and 7.5) had the highest malate titers (Fig. S2A). Second, buffer concentration was optimized. It was found out that 100 mM HEPES buffer (pH 7.5) was the best (Fig. S2B). Third, the optimal reaction temperature was 50 °C (Fig. S2C). Lastly, the effects of oxygen presence and the addition of a reducing agent DTT were investigated (Fig. S2D). The malate titer was enhanced from 3.04 mM under aerobic conditions to 4.21 mM under anaerobic conditions; 5.76 mM was obtained under aerobic conditions supplemented with DTT; the highest titer of 5.98 mM was obtained under anaerobic conditions with DTT. Finally, the optimized reaction conditions (i.e., 100 mM HEPES, pH 7.5, 50 °C, and anaerobic condition plus DTT) lead to 8.48 mM malate, 3.3 times that of 2.58 mM in the proof-of-concept reaction (Fig. S3). Because dissolved oxygen in the buffer had a negative effect on malate formation, an oxygen indicator resazurin was added into the reaction solution to indicate whether the dissolved oxygen was removed completely. Under aerobic conditions, the addition of DTT enabled to mitigate oxygen's negative effect with the decreased pink color, the malate titer was enhanced from 2.58 to 8.15 mM under aerobic conditions supplemented with DTT. De-oxidization by nitrogen flushing followed by an anaerobic chamber enabled to turn the pink color into colorless whether or not DTT was added. However, under anaerobic conditions, the addition of DTT enabled to promote the malate production from 4.72 mM to 8.48 mM (Fig. 4). These results suggested that

4. Discussion This study demonstrated that up to 52.4 mM malate was produced by the in vitro ATP-balanced pathway with 95.3% of the theoretical maximum yield through the complete utilization of starch (i.e., 27.5 mM glucose equivalent). It was notable that the combination of Modules 2.1 and 2.2 enabled in vitro ATP balance and regeneration without exogenous addition of ATP donors (Fig. 5). To improve ATP supplies, we chose the ATP-conservation pathway from starch to G3P (Module 1) instead of a typical glycolysis pathway, saving one ATP per glucose. Furthermore, in the CO2 fixation module from 3-PG or its derived metabolite to malate, three possible pathways were (1) ATPconsuming pathway (Module 3); (2) ATP-neutral pathway from PEP to OAA catalyzed by phosphoenolpyruvate carboxykinase, and (3) ATPneutral pathway from PEP to pyruvate catalyzed by pyruvate kinase (Zhang et al., 2009). The ATP-consuming enzymatic pathway was preferred due to its negative Gibbs free energy. The ATP-balanced pathway (Fig. 1C) had a self-balanced ATP supply and consumption without exogenously ATP donors. This in vitro pathway cannot be implemented by living microorganisms due to dual reasons: (1) maltodextrin, a polysaccharide, cannot be transported across cellular membrane and ATP-saving intermediate G1P production from starch cannot be assimilated by living cells; and (2) the overall ATP-neutral pathway cannot provide net ATP for basic maintenance and self157

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Fig. 4. Effects of reducing states on malate synthesis. The images with color change in the serum bottles (A) and the profiles of malate biosynthesis (B) under different reaction conditions. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

Fig. 5. Malate synthesis by the ATP-balanced systems with different ratios of ATP/ADP. The time courses for malate production (A) and the profiles of total turnover number (TTN) (B), where TTN = malate titer/(2*initial ATP addition).

Fig. 6. The profiles of titers and yields of malate by using different enzyme treatment for enhanced utilization of starch. The time course for malate production (A) by the 13-enzyme cocktail (open square), from IA-treated starch (solid circle), from IA-treated starch supplemented with 4GT (solid square), and from IA-treated starch supplemented with 4GT and PPGK (open circle), as well as malate yields by different enzyme cocktails (B). 158

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enzyme cocktails by enzyme immobilization (Sheldon, 2007; Sheldon and Woodley, 2017), use of thermophilic enzymes and enzyme engineering (Zhou et al., 2018), would be one of the future research directions. Enhanced enzyme stability could not only increase the economy of ivME but also enhance the net CO2 fixation yield in terms of the total substrate consumption that closed to the theoretical yield of the biotransformation step (i.e., 1 mol CO2 per mole malate). This study demonstrated a feasibility of CO2 fixation energized by starch for the production of malate. Although the final titer of malate was 52.4 mM from 27.5 mM glucose equivalent maltodextrin, lower than that of 253 mM from 178 mM glucose (Zhang et al., 2011) and 1.46 M from 2.72 M glycerol (Zambanini et al., 2016) by microbial fermentation, this enzyme cocktail had a higher product yield of 1.91 mol/mol glucose than those of 1.42 mol/mol glucose (Zhang et al., 2011) and 0.54 mol/mol glycerol (Zambanini et al., 2016). The in vitro reconstitution of the synthetic CO2 fixation biosystems featuring the ATP-balanced pathways could offer an alternative for the efficient conversion of CO2 into biochemicals. The platform does however still need to be further improved for future implementation in biomanufacturing, for example, by increasing cofactor stability by using lesscostly and more stable biomimics (Zachos et al., 2019) or the introduction of their salvage pathways (Opgenorth et al., 2014), by immobilizing enzymes for prolonged lifetime and easy separation of products and enzymes (Sheldon, 2007; Sheldon and Woodley (2017)), by implementing fast conversion at high substrate concentrations (Kim et al., 2018; You et al., 2017), by constructing multi-enzyme complexes featuring metabolite challenging to avoid labile metabolite degradation (Zhang, 2011), and so on.

duplication of cells. Anaerobic synthesis of succinic acid was one of the best examples of chemical energy supply for CO2 fixation, whereas PEPC was recognized as the primary enzyme for PEP carboxylation during succinate production in E. coli. Energetically, the PEPC-catalyzed reaction was strongly favored, but energy contained in PEP was lost in this reaction with the release of inorganic phosphate. Recruiting PEP carboxykinase (PCK) for PEP carboxylation could conserve ATP, the dominant route for carboxylation in these succinate-producing strains of E. coli and in succinate-producing bacteria (Zhang et al., 2009). Anoxic conditions and/or the addition of a reducing agent DTT were important for in vitro malate synthesis (Fig. 4B). The mechanism here was different from that of anaerobic xylose-to-inositol biotransformation mainly due to that Maillard reaction (Cheng et al., 2019) because the reaction temperature here was not very high, there were no reducing sugars (i.e., xylose) and no brown color was observed. The oxygenic conditions were thought to deactivate some enzymes that require sulfhydryl groups for their activities. For example, molecular oxygen oxidizes the essential active-center cysteine residue Cys-149 of GAPDH to a sulfenic acid and not to a disulfide, leading to the conversion of enzyme from a dehydrogenase to an acyl phosphatase (Allison and Benitez, 1972). This deactivation can be mitigated by adding reducing agents and their stability and recyclability can also be increased significantly under anoxic conditions (Crans and Whitesides, 1985). In addition, some intermediate metabolites could undergo oxidation reactions, such as the oxidation of DHAP to hydroxypyruvaldehyde phosphate (Hixon et al., 1996). Furthermore, it was known that molecular oxygen under aerobic conditions can oxidize the nicotinamide ring of NADH at evaluated temperatures, leading to the fast decomposition of NADH (Hofmann et al., 2010). Therefore, it was essentially important to keep NADH stable at 50 °C by providing the anaerobic conditions and/or the addition of DTT. When aerobic conditions were used, the re-addition of NADH was essential for continuous production of lactate by the enzyme cocktail at 50 °C (Ye et al., 2012). Our previous results suggested that NADH with reducing agents was pretty stable for 24 h under anaerobic conditions even at 80 °C (Kim et al., 2018). Thermal degradation of labile metabolites in vitro, such as G3P and DHAP, could lead to the loss of free energy and carbon, resulting in the accumulation of dead-end compounds (Kouril et al., 2012). It is known that some intermediates, such as G3P, DHAP and 1,3-BPG, have half-life times of the order of minutes at 80 °C (Kouril et al., 2013). Living thermophilic cells have developed a few strategies to address rapid degradation of labile metabolites, such as macromolecular crowding, enzyme complexes featuring metabolite channeling, highly reducing environment, or the use of pathways without thermolabile intermediates (Daniel and Cowan, 2000). The results of increased malate yields with increasing enzyme loadings (Fig. S5) implied that high enzyme loading like macromolecular crowding effects could avoid degradation of labile metabolites. Also, higher enzyme loadings could enhance their stability (Myung and Zhang, 2013). The optimal reaction temperature of 50 °C was chosen as a tradeoff between the thermostability of intermediates and the enzyme activities (Fig. S2C). One of further improvements of in vitro biosystems could be the construction of enzyme complexes facilitating fast transfer of labile metabolites among cascade enzymes, like previous efforts in the production of sweet hydrogen (Kim et al., 2018). Although ivME demonstrated very high yields of malate (Fig. 6), a significant amount of CO2 was released during the fermentation of E. coli for the production of thermophilic enzymes. When two steps of enzyme fermentation featuring CO2 emissions and malate biotransformation featuring CO2 fixation were combined, the net yield of CO2 fixation was not as high as its theoretical yield of the biotransformation step. If the lifetimes of enzymes cocktails (i.e., total turn-over times of enzymes) were improved to the level of an industrial enzyme – immobilized glucose isomerase, the ratio of CO2 emissions to fixation would be very small. Therefore, prolonging the lifetimes of

5. Conclusions It is attractive to develop innovative strategies to convert CO2 into multi-carbon biocommodities. Here we designed the in vitro artificial ATP-balanced pathway for CO2 fixation and malate synthesis energized by starch. Up to 52.4 mM malate was obtained with 95.3% of the theoretical maximum yield (1.91 mol/mol glucose). This ATP-balanced pathway design enables the system to automatically adjust its fluxes between Modules 2.1 and 2.2 regardless of ATP/ADP ratios. Anaerobic reaction conditions and/or the addition of reducing agent DTT were of importance for creating a reductive environment for proper working of the enzyme cocktail and minimizing degradation of intermediates. Acknowledgment This work was supported by the National Natural Science Foundation of China (Grant No. 31700033). Appendix A. Supplementary data Supplementary data related to this article can be found at https:// doi.org/10.1016/j.ymben.2019.07.005. References Ajjawi, I., Verruto, J., Aqui, M., Soriaga, L.B., Coppersmith, J., Kwok, K., Peach, L., Orchard, E., Kalb, R., Xu, W., others, 2017. Lipid production in Nannochloropsis gaditana is doubled by decreasing expression of a single transcriptional regulator. Nat. Biotechnol. 35 (7), 647–652. Allison, W.S., Benitez, L.V., 1972. An adenosine triphosphate-phosphate exchange catalyzed by a soluble enzyme couple inhibited by uncouplers of oxidative phosphorylation. Proc. Natl. Acad. Sci. U.S.A. 69 (10), 3004–3008. Angermayr, S.A., Paszota, M., Hellingwerf, K.J., 2012. Engineering a cyanobacterial cell factory for production of lactic acid. Appl. Environ. Microbiol. 78 (19), 7098–7106. Ardao, I., Zeng, A.-P., 2013. In silico evaluation of a complex multi-enzymatic system using one-pot and modular approaches: application to the high-yield production of hydrogen from a synthetic metabolic pathway. Chem. Eng. Sci. 87, 183–193. Beer, B., Pick, A., Sieber, V., 2017. In vitro metabolic engineering for the production of alpha-ketoglutarate. Metab. Eng. 40, 5–13. Cheng, K., Zhang, F., Sun, F., Chen, H., Zhang, Y.-H.P., 2015. Doubling power output of

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