- Open Access
Engineering a cyanobacterium as the catalyst for the photosynthetic conversion of CO2 to 1,2-propanediol
© Li and Liao; licensee BioMed Central Ltd. 2013
Received: 14 September 2012
Accepted: 7 January 2013
Published: 22 January 2013
The modern society primarily relies on petroleum and natural gas for the production of fuels and chemicals. One of the major commodity chemicals 1,2-propanediol (1,2-PDO), which has an annual production of more than 0.5 million tons in the United States, is currently produced by chemical processes from petroleum derived propylene oxide, which is energy intensive and not sustainable. In this study, we sought to achieve photosynthetic production of 1,2-PDO from CO2 using a genetically engineered cyanobacterium Synechococcus elongatus PCC 7942. Compared to the previously reported biological 1,2-PDO production processes which used sugar or glycerol as the substrates, direct chemical production from CO2 in photosynthetic organisms recycles the atmospheric CO2 and will not compete with food crops for arable land.
In this study, we reported photosynthetic production of 1,2-PDO from CO2 using a genetically engineered cyanobacterium Synechococcus elongatus PCC 7942. Introduction of the genes encoding methylglyoxal synthase (mgsA), glycerol dehydrogenase (gldA), and aldehyde reductase (yqhD) resulted in the production of ~22mg/L 1,2-PDO from CO2. However, a comparable amount of the pathway intermediate acetol was also produced, especially during the stationary phase. The production of 1,2-PDO requires a robust input of reducing equivalents from cellular metabolism. To take advantage of cyanobacteria’s NADPH pool, the synthetic pathway of 1,2-PDO was engineered to be NADPH-dependent by exploiting the NADPH-specific secondary alcohol dehydrogenases which have not been reported for 1,2-PDO production previously. This optimization strategy resulted in the production of ~150mg/L 1,2-PDO and minimized the accumulation of the incomplete reduction product, acetol.
This work demonstrated that cyanobacteria can be engineered as a catalyst for the photosynthetic conversion of CO2 to 1,2-PDO. This work also characterized two NADPH-dependent sADHs for their catalytic capacity in 1,2-PDO formation, and suggested that they may be useful tools for renewable production of reduced chemicals in photosynthetic organisms.
Many natural metabolites containing bi-functional groups such as succinate, lactate, and 3-hydroxybutaoate can be used as monomers to make polymers, and have long been produced biologically by fermentation processes. However, production of diols from renewable source represents unique challenges partially because they are not typical fermentation products and that they are more reduced compared to the average carbon redox state in biological systems . Over the past decade, progress in synthetic biology and metabolic engineering have enabled substantial achievements in diol production [1–4], mostly from sugars, glycerol, or biomass feedstocks. Direct production of chemicals from CO2 in photosynthetic organisms [5–7] and lithoautotrophic organisms [8–10] have been proposed to be advantageous in particular situations. This work aims to produce 1,2-propanediol (1,2-PDO) directly from CO2 by an engineered cyanobacterium, Synechococcus elongatus PCC 7942.
Results and discussion
Designing of the 1,2-PDO production pathway
In light conditions, cyanobacteria fix CO2 via the Calvin-Benson-Bassham (CBB) cycle which is powered by ATP and NADPH generated by the photosystems (Figure 1A). Two CBB cycle intermediates, fructose-6-phosphate (F6P) and glyceraldehydes-3-phosphate (GAP), serve as the branch points of carbon leaving the CBB cycle to the central metabolism for glycogen synthesis and glycolysis, respectively. While glycogen synthesis is the major carbon and energy storage pathway, glycolysis and TCA cycle produce building blocks for cell growth. The synthesis of 1,2-PDO, on the other hand, starts from another CBB cycle intermediate, dihydroxyacetonephosphate (DHAP). The introduction of one extra branch point can potentially increase the flux of output carbon from the CBB cycle, which has been suggested to be beneficial for increasing photosynthesis efficiency in higher plants [18, 19], but may also disrupt the normal flux distribution in the cell.
To synthesize 1,2-PDO, DHAP is first converted to methylglyoxal (Figure 1A) by methylglyoxal synthase (encoded by mgsA in E. coli). Methyglyoxal is very toxic to the cells  and needs to be efficiently utilized by downstream enzymes. Two different metabolic routes have been shown to synthesize 1,2-PDO from methyglyoxal (Figure 1B) . The first involves reduction of methyglyoxal by the glycerol dehydrogenase (encoded by gldA in E. coli) to lactaldehyde, which is further reduced by the 1,2-propanediol reductase (encoded by fucO in E.coli) to yield the final product. The second route includes an alcohol dehydrogenase (such as the broad-substrate range aldehyde reductase encoded by yqhD in E. coli) to produce acetol as the intermediate, which is then converted to 1,2-PDO by gldA. The latter route was chosen to introduce into S. elongatus because yqhD gene has been previously overexpressed in this organism for biofuel production and showed relatively good performance, possibly due to its NADPH-specific cofactor preference.
Introduction of the 1,2-PDO biosynthesis genes
Production of 1,2-PDO
If NADH is really the limiting factor in our production scenario, the partially reduced intermediate acetol may accumulate. In fact, at the end of the production, around 16mg/L acetol was accumulated, which was comparable to the level of 1,2-PDO (~22mg/L) (Figure 3D). In addition, acetol was only detected after 4 days and kept accumulating during the late stage of production (data not shown). These results are consistent with the above-mentioned hypothesis and suggest that the NADH-dependent reduction of acetol catalyzed by gldA might be the limiting step in the 1,2-PDO production pathway.
Improving 1,2-PDO production using NADPH-dependent secondary alcohol dehydrogenases
To overcome the bottleneck of the 1,2-PDO production in LH21, one possible strategy is to overexpress the soluble transdehydrogenase (STH) which produces NADH at the expense of NADPH. However, genes encoding this enzyme have not been found in S. elongatus genome. Heterologous overexpression of the Pseudomonas aeruginosasth gene in cyanobacteria has been shown to be instable and caused growth defect .
Kinetics parameters of gldA and secondary alcohol dehydrogenases (sADH) for acetol
K m (mM)
K cat (S-1)
K cat /K m (mM-1S-1)
C. beijerinckii and T. brockii adh were cloned and introduced into the cyanobacterial genome to replace gldA. The resulting strains are named LH22 and LH23, respectively (Figure 2A). RT-PCR was also performed to verify the expression of these genes (Figure 2B, C). Enzyme assays with crude cell extract of LH22 and LH23 further verified that both C. beijerinckii and T. brockii sADH were functionally overexpressed and showed higher activities of NAD(P)H-dependent acetol reduction compared to that in LH21. Especially, the C. beijerinckii sADH overexpression in LH22 delivered the highest activity (Figure 3B, C).
Production using strains LH22 and LH23 yielded significantly higher 1,2-PDO titer (~150 and 80mg/L, respectively) compared to that of LH21 (Figure 4B). Notably, the high production rate was maintained through the 10 days of production. In consistent with the hypothesis mentioned in the previous section, the high level of NADPH-dependent acetol reduction activity in LH22 and LH23 also significantly reduced the accumulation of the intermediate acetol (Figure 3D).
Despite its great significance to metabolic engineers, the information on intracellular NAD(P)H level during different growth phases and growth conditions in cyanobacteria is very limited. Although it is believed that NADPH is more abundant than NADH in cyanobacteria , only a few studies discussed its role in biofuel/biochemical synthesis from CO2[6, 22]. The science behind efficient conversion of CO2 to chemicals and fuels is still in its infancy and the NADPH driving force theory still needs to be extensively tested, which requires the accumulation of empirical evidence in more production scenario, as well as fundamental studies on NAD(P)H levels and their regulation. In our case, other factors may also contribute to the difference between the production levels of the NADH and NADPH-dependent pathways. For example, the NADPH-dependent enzymes may be better folded and more active when expressed in cyanobacteria. And different level of physiological fitness may be caused by overespression of different enzymes, although all production strains showed the same growth phenotype as the wildtype.
In this work, we demonstrated the 1,2-PDO production from CO2 for the first time by the engineered cyanobacterium S. elongatus PCC 7942. By exploiting sADHs which have not been reported for 1,2-PDO production previously, a completely NADPH dependent pathway was built to channel the CBB cycle intermediate DHAP for 1,2-PDO production without accumulating the pathway intermediate, acetol. The best strain LH22, which harbors mgsA and yqhD both from E. coli and the adh from C. beijerinckii, produced ~150mg/L 1,2-PDO.
This work revealed the great potential of the vast NADPH pool in photosynthetic cyanobacteria as a robust driving force for the production of chemicals. Among the chemicals that have been produced biologically in industrial scale, a significant number of them are synthesized by NADPH consuming pathways. For example, in amino acid production, studies have shown that increasing the NADPH pool can improve the production performance [26, 27]. However, in most of the heterotrophic microorganisms, NADPH is mainly generated through the pentose phosphate pathway and TCA cycle and its pool size is relatively small compared to that of the NADH. On the other hand, photosynthetic organisms maintain high intracellular NADPH level. The unique metabolic feature of photosynthetic organisms provides great opportunities for the production of chemicals through NADPH dependent pathways.
Chemicals and reagents
All chemicals were purchased from Sigma-Aldrich (St. Louis, MO) or Fisher Scientifics (Pittsburgh, PA). Restriction enzymes were purchased from New England BioLabs (Ipswich, MA). The Rapid DNA ligation kit was from Roche (Mannheim, Germany). KOD DNA polymerase was from EMD Chemicals (San Diego, CA). Oligonucleotides were purchased from IDT (San Diego, CA).
Medium and culture condition
All S. elongatus PCC 7942 strains were grown on BG-11 medium (Sigma-Aldrich) containing 50mM NaHCO3 in shake flasks. Plates contain 1.5% (w/v) agar. For 1,2-PDO production, 50mL culture was grown in 250mL shake flasks under 100 μE/s/m2 light supplied by four Lumichrome F30W-1XX 6500K 98CRI light tubes, at 30°C. Cell growth was monitored by measuring OD730. 1mM IPTG was added to induce the gene expression at OD730 of around 1. Daily, samples were taken for analysis and 50mM NaHCO3 was added. IPTG concentration in the culture was maintained to 1mM by adding appropriate amount of fresh IPTG to compensate the IPTG lost from sampling. Spectinomycin was added for LH21, LH22, and LH23 at a final concentration of 20mg/L. E. coli strains were grown in LB medium. And a spectinomycin concentration of 50mg/L was used where appropriate.
Plasmids used in this study
Used for strain
NSI targeting vector
Contain C. beijerinckii adh gene codon optimized for E.coli
Contain T. brockii adh gene codon optimized for E.coli
NSI targeting. LacIq; Ptrc::gldA, yqhD, mgsA; Spec R
NSI targeting. LacIq; Ptrc:: C.beijerinckii adh, yqhD, mgsA; Spec R
NSI targeting. LacIq; Ptrc::T.brockii adh, yqhD, mgsA; Spec R
E.coli vector whose backbone was used to build his tag protein expression plasmids used in this study.
Vector for N-terminal 6Xhis-tagged protein expression
The pZElac inserted with the T5 promoter/lacO::6Xhis part from pQE-9.
E.coli vector for his tag- E.coli gldA expression.
E.coli vector for his tag- C. beijerinckii adh expression.
E.coli vector for his tag- T. brockii adh expression.
Primers used in this study
Used for plasmid
pQE XhoI fwd
pQE Acc65I rev
his ad up rev
his ad down fwd
his ad up_gldA fwd
his ad down_gldA rev
his ad up_CB fwd
his ad down_CB rev
his ad up_TB fwd
his ad down_TB rev
Briefly, to construct plasmid GYM, gldA, mgsA, and yqhD were amplified from E. coli genomic DNA using primer pairs gldA SpeI fwd/gldA_YqhD rev, gldA_YqhD fwd/YqhD_mgsA rev, and YqhD_mgsA fwd/mgsA NotI rev, respectively. The PCR products were purified and linked into an artificial operon using Splicing by overhang extension (SOE) PCR using primers gldA SpeI fwd/mgsA NotI rev. The PCR product was digested with restriction enzymes SpeI and NotI and then inserted into the NSI targeting vector pAM2991. The CYM and TYM plasmids were constructed similarly. The C. beijerinckii adh and T. brockii adh genes were amplified from plasmids pZE12-alsS-alsD-CBADH and pZE12-alsS-alsD-TBADH , respectively, using primer pairs CBSADH SpeI fwd/CBSADH_yqhD rev and TBSADH SpeI fwd/TBSADH_yqhD rev, respectively. In order to amplify yqhD gene that have overlapping region with the C. beijerinckii adh and T. brockii adh, the forward primer for yqhD amplification was CBSADH_yqhD fwd and TBSADH_yqhD fwd, respectively.
To purify the 6xHis-tagged gldA and secondary alcohol dehydrogenases, plasmids his-gldA, his-CB, and his-TB were constructed. Briefly, the T5 promoter/lacO and 6xHis tag fragment of pQE-9 (Qiagen) was amplified using primers pQE XhoI fwd and pQE Acc65I rev and then digested and inserted at XhoI/Acc65I sites of the plasmid pZElac . The resulted plasmid was named pZElac-his. To insert the E. coli gldA, C. beijerinckii adh and T. brockii adh genes in pZElac-his, the isothermal DNA assembly method  was used. The primers his ad up rev and his ad down fwd were used to amplify the vector backbone using pZElac-his as template. And the primer pairs his ad up_gldA fwd/his ad down_gldA rev, his ad up_CB fwd/his ad down_CB rev, and his ad up_TB fwd/his ad down_TB rev were used to amplify the corresponding genes. The gene amplification products were assembled with the backbone.
Protein purification and enzyme kinetics study
The plasmid his-gldA, his-CB, and his-TB were transformed into BL21 cells. The transformants were cultured in 40 mL LB medium containing 100mg/L ampicillin. After the cells reached mid-log phase, 1mM IPTG was added to induce protein expression followed by incubation at 30°C overnight. The cells were collected by centrifugation and the recombinant proteins were purified using His-Spin Protein Miniprep kit (Zymo research Corporation, CA) according to the manufacturer’s instructions. The purified proteins were checked by SDS-PAGE for homogeneity and quantified by Bradford assay (Bio-Rad, Hercules, CA).
Dehydrogenase activity was measured by monitoring the absorbance decrease of NADH or NADPH at wavelength of 340 nm. To determine the kinetic parameters, the assay reaction was prepared with Tris-HCl buffer (50 mM, pH = 7.5) containing 200 μM of NADH or NADPH and various concentration of acetol ranging from 0.05 mM to 20 mM at room temperature. The K m and K cat values were obtained by non-linear fitting with the Michaelis–Menten equation.
Transformation and selection
S. elongatus PCC7942 cells transformed as described . The transformed cells were spread on BG-11 plates with 20mg/ml spectinomycin and incubated in light to select for recombinants. Colonies were verified by PCR and inoculated into liquid BG-11 medium with 20 mg/ml spectinomycin for further tests.
For sADH enzyme assays, different cyanobacterial strains were grown in 30mL BG-11 medium in 125mL shake flask under light condition and induced with 1mM IPTG at the OD730 of around 1. After overnight induction, cells were harvested and resuspended in 1mL Buffer A (100mM Tris-HCl, pH = 8.0). Cell lysates were prepared by bead beating followed by centrifugation at 10,000 × g for 20min at 4°C. Cell. 10μL cell lysate was used in 200μL reaction system which also contained 100mM Tris-HCl, pH = 8.0, 200μL NAD(P)H, and 20mM acetol (or no substrate for negative control). The reaction was started by adding the substrate, and the OD340 was monitored. The soluble protein concentrations in cell lysates were quantified using Quick Start Bradford Protein Assay (Bio-Rad, CA) according to manufacturer’s instructions. Similar method was used to determine the activity of methylglyoxal reduction.
The activity of methylglyoxal synthase was determined by the previously reported method . Briefly, the reaction mixture at 30°C contained, in 0.5 ml: Tris-HCl buffer, pH7.5 (50mM), dihydroxyacetone phosphate (20mM) and cell lysate. The reaction was allowed to proceed for 10min. The methylglyoxal formed was measured colorimetrically by taking 0.1 ml samples into 0.33 ml of 2,4-dinitrophenylhydrazine reagent (0.1% 2,4-dinitrophenylhydrazine in 2M-HCI) plus 0.9ml of water. After incubation at 30°C for 15min, 1.67ml of 2.5M NaOH was added and the OD555 measured after a further 15min. A molar extinction coefficient of 4.48 × 104 was used to convert the readings into nmol of methylglyoxal.
For RT-PCR enzyme assays, different cyanobacterial strains were grown in 30mL BG-11 medium in 125mL shake flask under light condition and induced with 1mM IPTG at the OD730 of around 1. After overnight induction, total RNA was extracted using RiboPure-Bacteria Kit (Life Technologies, NY). RNA was quantified using Nanodrop. After treatment with the TURBO DNA-free kit (Life Technologies, NY), cDNA was synthesized using iScript cDNA Synthesis kit (Bio-Rad, CA). PCR was performed using the specific primers listed in Table 2. The PCR product was then checked by electrophoresis on 2% agarose gel and stained with ethidium bromide.
Quantification of 1,2-PDO and acetol
1mL samples were taken daily from production culture. After centrifugation, the supernatant was taken for 1,2-PDO and acetol analysis by Agilent 1200 highpressure liquid chromatography (HPLC) system equipped with an autosampler (Agilent Technologies),a Bio-Rad (Bio-Rad Laboratories, Hercules, CA) Aminex HPX87 column (5 mM H2SO4, 0.6 mL/min, column temperature at 35°C) and a refractive index detector (RID) module.
We thank Dr. Yajun Yan for helpful discussion and the plasmids pZE12-alsS-alsD-CBADH and pZE12-alsS-alsD-TBADH. We thank Dr. Wendy Higashide for assistance on experiments.
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