NOVEL GENETICALLY ENGINEERED MICROORGANISM CAPABLE OF GROWING ON FORMATE, METHANOL, METHANE OR CO2

Abstract

The present invention relates to a genetically engineered microorganism expressing (i) formate tetrahydrofolate (THF) ligase, methenyi-THF cyclohydrolase and methylene-THF dehydrogenase, (ii) the enzymes of the glycine cleavage system (GCS), (iii) serine deaminase and serine hydroxymethyltransferase (SHMT), (iv) an enzyme increasing the availability of NADPH, and (v) optionally formate dehydrogenase (FDH), and wherein the genetically engineered microorganism has been genetically engineered to express at least one of the enzymes of (i) to (v), wheren said enzyme is not expressed by the corresponding microorganism that has been used to prepare the genetically engineered microorganism, and wherein the enzymes of (i) to (v) are genomically expressed.

Claims

1. A genetically engineered microorganism expressing (i) formate tetrahydrofolate (THF) ligase, methenyl-THF cyclohydrolase and methylene-THF dehydrogenase, (ii) the enzymes of the glycine cleavage system (GCS), (iii) serine deaminase and serine hydroxymethyltransferase (SHMT), (iv) an enzyme increasing the availability of NADPH, and (v) optionally formate dehydrogenase (FDH), and wherein the genetically engineered microorganism has been genetically engineered to express at least one of the enzymes of (i) to (v), wheren said enzyme is not expressed by the corresponding microorganism that has been used to prepare the genetically engineered microorganism, and wherein the enzymes of (i) to (v) are genomically expressed.

2. The genetically engineered microorganism of claim 1, wherein the enzymes of (i) to (v) are expressed from genomic safe spots.

3. The genetically engineered microorganism of claim 1 or 2, wherein one or more of the enzymes of (i) to (v) are expressed under the control of a strong constitutive promoter and/or a modified ribosome binding site.

4. The genetically engineered microorganism of any one of claims 1 to 3, wherein the enzyme of (iv) is at least 2-fold, preferably at-least 3-fold, more preferably at least 4-fold and most preferably at least 5-fold higher expressed than the enzymes of (i) to (iii).

5. The genetically engineered microorganism of any one of claims 1 to 4, wherein the enzyme increasing the availability of NADPH is membrane transhydrogenase (PntAB), glucose 6-phosphate dehydrogenase (Zwf), 6-phosphogluconate dehydrogenase (Gnd), or malic B enzyme (MaeB), isocitrate dehydrogenase (lcd), and is preferably PntAB.

6. The genetically engineered microorganism of claim 5, wherein an overexpression of PntAB is achieved by introducing a mutation into the promoter region of pntAB, wherein the mutation of pntAB is preferably a single-base pair substitution in the promoter region of pntAB.

7. The genetically engineered microorganism of any one of claims 1 to 6, wherein an overexpression of FDH is at least partly achieved by introducing a mutation into the 5′ untranslated region of FDH, wherein the mutation of FDH is preferably a single-base pair substitution in the 5′ untranslated region of FDH.

8. The genetically engineered microorganism of any one of claims 1 to 7, wherein the microorganism is auxotrophic for serine, glycine and C.sub.1 moieties.

9. The genetically engineered microorganism of any one of claims 1 to 8, wherein the microorganism is a bacterium, preferably a proteobacterium, more preferably an enterobacterium and most preferably E. coli.

10. The genetically engineered microorganism of any one of claims 1 to 9, wherein the microorganism is capable of converting methanol to formate.

11. The genetically engineered microorganism of any one of claims 1 to 9, wherein the microorganism is capable of converting methane to formate.

12. The genetically engineered microorganism of any one of claims 1 to 9, wherein the microorganism is capable of converting CO.sub.2 to formate.

13. A method for growing the microorganism of any one of claims 1 to 9 which expresses FDH, comprising culturing the microorganism under growth conditions comprising formate as the sole carbon source.

14. A method for growing the microorganism of claim 10, comprising contacting the microorganism under growth conditions comprising methanol as the sole carbon source.

15. A method for growing the microorganism of claim 11 which expresses FDH, comprising culturing the microorganism under growth conditions comprising methane as the sole carbon source.

16. A method for growing the microorganism of claim 12 which expresses FDH, comprising culturing the microorganism under growth conditions with CO.sub.2 as the sole carbon source.

Description

[0104] The figures show.

[0105] FIG. 1—The synthetic reductive glycine pathway is similar in structure to the reductive acetyl-CoA pathway. Yet, while the latter pathway is restricted to anaerobic conditions, the former can operate under aerobic conditions. Both pathways are highly ATP-efficient, as only 1-2 ATP molecules are consumed in the conversion of formate to pyruvate (e.g., instead of 7 by the Calvin Cycle). Molecular structure in brown corresponds to a sub-structure of tetrahydrofolate. Enzymes of the reductive glycine pathway, as implemented in this study, are indicated in purple (Lpd, unlike the other enzymes of the glycine cleavage system, was not overexpressed). ‘Me’ corresponds to Methylobacterium extorquens and ‘Ec’ corresponds to Escherichia coli. Division of the pathway into modules, as explained in the text, is shown in light brown to the right of the figure.

[0106] FIG. 2—Modular establishment of the reductive glycine pathway. (A) Selection scheme of C.sub.1M and C.sub.2M for the biosynthesis of C.sub.1-moieties, glycine, and serine. (B) Overexpression of C.sub.1M and C.sub.2M enabled growth with formate (and CO.sub.2) as sole source of C.sub.1-moieties, glycine, and serine. (C) Selection scheme of C.sub.1M, C.sub.2M, and C.sub.3M to generate biomass building blocks, where acetate oxidation provides reducing power and energy. Deletion of aceA prevents acetate from being used as a carbon source. (D) Overexpression of C.sub.1M, C.sub.2M, and C.sub.3M enabled growth with formate as source of biomass and acetate as an energy source. Genomic integration of C.sub.3M was performed in strain in which the endogenous glyA and sdaA were deleted. (E) Selection scheme of C.sub.1M, C.sub.2M, C.sub.3M, and EM to use formate as sole carbon and energy source. (F) Growth on formate is demonstrated only when all four modules are overexpressed. Genomic overexpression is indicated by ‘g’, while overexpression from a plasmid is indicated by ‘p’. Experiments were conducted at 10% CO.sub.2 within 96-well plates and were performed in triplicate, which displayed identical growth curves (±5%), and hence were averaged. Doubling times (DT) shown in the figure.

[0107] FIG. 3—Short term evolution improves growth on formate. (A) Test-tube cultivation on formate as sole carbon source. The vertical small red arrows correspond to the addition of formate, increasing the concentration in the medium by 30 mM. Upon reaching an OD.sub.600 of 0.4, cells were reinoculated into a new test-tube with an initial OD.sub.600 of 0.03-0.05. Error bars correspond to standard deviation of 2 experiments. 6 exemplifying cycles of cultivation are shown. (B) Doubling time decreased with cultivation cycle. Error bars correspond to standard deviation of 2 experiments. (C) Growth of the evolved strain (in test-tube) is directly coupled to a decrease in formate concentration. Error bars correspond to standard deviation of 2 experiments. (D) Cultivation of the evolved strain on formate as a sole carbon source within a 96-well plate. Experiments were conducted at 10% CO.sub.2. Plate reader experiments were performed in triplicate, which displayed identical growth curves (±5%), and hence were averaged. Doubling times (DT) are shown in the figure. DT were considerably shorter in the plate reader than in test-tube as the measurements in were more accurate (taken every 10 minutes rather than once per day) and since the conditions are different (e.g., more stable cultivation environment in the plate reader).

[0108] FIG. 4—Labeling pattern of proteinogenic amino acids confirms the activity of the reductive glycine pathway. As elaborated in FIG. 13, the labeling pattern is consistent with the assimilation of formate and CO.sub.2 via the synthetic pathway, and indicates low cyclic flux via the TCA cycle. Numbers written in italics above the bars correspond to the overall fraction of labeled carbons.

[0109] FIG. 5—Engineered growth on methanol. (A) Methanol can be assimilated via the activity of methanol dehydrogenase (MDH), where formaldehyde is oxidized to formate via the native activity of the glutathione system. (B) The Methanol Module (MM) converts methanol to formate and provides the cell with reducing power and energy. (C) Overexpression of MDH from Bacillus stearothermophilus (BsMDH) within the gC.sub.1M gC.sub.2M gC.sub.3M gEM strain, carrying a mutation in the promoter of the pntAB operon (FIG. 12), enabled growth on methanol within a 96-well plate. Experiments were conducted at 10% CO.sub.2. Plate reader experiments were performed in triplicate, which displayed identical growth curves (±5%), and hence were averaged. (D) Comparison of growth on methanol (shown are final cell densities) with different expressed enzymes and at different genetic backgrounds. NAD-dependent MDH from several organisms was tested: Bacillus stearothermophilus (BsMDH), Corynebacterium glutamicum (CgMDH), and Cupriavidus necator N-1 (CnMDH), as well as two MDHs from B. methanolicus (BmMDH2 and BmMDH3) and an improved variant (BmMDH2*, carrying Q5L A363L modifications). Formaldehyde dehydrogenases from Pseudomonas putida (PpFADH; SEQ ID NOs: 49 and 50) and Pseudomonas aeruginosa (PaFADH; SEQ ID NOs: 51 and 52) was further tested. (E) Labeling pattern of proteinogenic amino acids upon feeding with .sup.13C-methanol/.sup.12-CO.sub.2 is identical that with .sup.13C-formate/.sup.12-CO.sub.2 (FIG. 4), confirming the activity of the reductive glycine pathway. Numbers written in italics above the bars correspond to the overall fraction of labeled carbons.

[0110] FIG. 6—Schematic overview of overexpression strategy. Gene overexpression from plasmid is shown in the left column while genomic overexpression is shown in the right column.

[0111] Promoter and ribosome binding sites are as described in a previous manuscript*. Genomic ‘safe spots’ were described previously**. (* S. Wenk, O. Yishai, S. N. Lindner, A. Bar-Even, An engineering approach for rewiring microbial metabolism. Methods Enzymol 608, 329-367 (2018); ** M. C. Bassalo et al., Rapid and efficient one-step metabolic pathway integration in E. coli, ACS synthetic biology 5, 561-568 (2016))

[0112] FIG. 7—Replacement of the native promoter of the GCV operon with a strong constitutive promoter increases gene expression 20-50 fold in a serine auxotroph (SerAux) strain (ΔserA Δkbl ΔltaE ΔaceA). Transcript levels were normalized to the expression of the rrsA gene and are shown relative to the expression of a WT (non-serine auxotroph) strain. As a comparison, the transcript levels induced by a weak constitutive promoter and moderate constitutive promoter are shown*. Experiments were performed in triplicate. (* S. Wenk, O. Yishai, S. N.

[0113] Lindner, A. Bar-Even, An engineering approach for rewiring microbial metabolism. Methods Enzymol 608, 329-367 (2018))

[0114] FIG. 8—Different expression approaches of the genes of C3M -glyA and sdaA—affect growth via the reductive glycine pathway, with acetate serving as an energy source. Expression on a plasmid resulted in an identical growth regardless of the promoter strength (green and blue lines). Overexpression of sdaA alone failed to achieve growth (pink and purple lines). Genomic expression (after deletion of endogenous glyA and sdaA) resulted in better growth when gene expression was controlled by a medium strength ribosome binding site (‘C’, pale blue line) than by a strong ribosome binding site (‘A’, brown line). ‘g’ corresponds to genomic expression and ‘p’ to expression on a plasmid. Origin and replication, promoters, and ribosome binding sites are described in a previous study*. (* S. Wenk, O. Yishai, S. N. Lindner, A. Bar-Even, An engineering approach for rewiring microbial metabolism. Methods Enzymol 608, 329-367 (2018))

[0115] FIG. 9—Different expression approaches of the genes of EM—formate dehydrogenase—affect growth via the reductive glycine pathway. Expression on a plasmid supported growth. Genomic overexpression supported growth only when the ribosome binding site was of the highest strength (‘A’). ‘g’ corresponds to genomic expression and ‘p’ to expression on a plasmid. Origin and replication, promoters, and ribosome binding sites are described in a previous study.* (* S. Wenk, O. Yishai, S. N. Lindner, A. Bar-Even, An engineering approach for rewiring microbial metabolism. Methods Enzymol 608, 329-367 (2018))

[0116] FIG. 10—Number of colony forming units increases monotonically with OD600 for cells growing on formate as sole carbon source.

[0117] FIG. 11—Cell growth on formate directly correlates with increased medium pH due to the accumulation of OH.sup.−.

[0118] FIG. 12—Two mutations emerged within the formatotrophic strain after a short period of evolution. (A) A point mutation in the 5′-UTR of the FDH gene. (B) A point mutation in the promoter of the pntAB gene. Strain K4 corresponds to a strain in which the four modules of the reductive glycine pathway were introduced into its genome, that is, gC1M gC2M gC3M gEM, while strain K4e the same strain after short term evolution.

[0119] FIG. 13—Change in transcript level in the evolved strain. (A) Levels of FDH transcript increased 2.7-fold in the evolved strain. (B) Levels of pntAB transcript increased by ˜14-fold in the evolved strain. In both cases transcript levels were normalized to the rrsA gene and are shown relative to the expression within a nonevolved strain. Experiments were performed in triplicate. Strain K4 corresponds to a strain in which the four modules of the reductive glycine pathway were introduced into its genome, that is, gC1M gC2M gC3M gEM, while strain K4e the same strain after short term evolution.

[0120] FIG. 14—Evolved strain displays 7.4-fold higher activity of FDH in cell extract. FDH activity was measured in 96-well plate by the addition of formate and NAD+ and was followed by increase in absorbance at 340 nm by the accumulation of NADH. The results were normalized to mg of total cell protein. Strain K4 corresponds to a strain in which the four modules of the reductive glycine pathway were introduced into its genome, that is, gC1M gC2M gC3M gEM, while strain K4e the same strain after short term evolution.

[0121] FIG. 15—Introduction of the two mutations found in genome sequencing of the evolved strain (5′UTR of fdh and promoter region of pntAB) improved growth on formate dramatically and resulted in a growth pattern very similar to that of the evolve strain (see FIG. 3C). Cultivation of the evolved strain on formate as a sole carbon source within a 96-well plate. Experiments were conducted at 10% CO2. Plate reader experiments were performed in triplicate, which displayed identical growth curves (±5%), and hence were averaged. Strain K4 corresponds to a strain in which the four modules of the reductive glycine pathway were introduced into its genome, that is, gC1M gC2M gC3M gEM.

[0122] FIG. 16—Addition of 100 mM sodium bicarbonate enables growth on higher concentrations for formate, as demonstrated with the evolved K4 strain and a K4 strain to the genome of which the two mutations found in the evolved strain were introduced. Cultivation of the evolved strain on formate as a sole carbon source within a 96-well plate. Experiments were conducted at 10% CO2. Plate reader experiments were performed in triplicate, which displayed identical growth curves (±5%), and hence were averaged. Strain K4 corresponds to a strain in which the four modules of the reductive glycine pathway were introduced into its genome, that is, gC1M gC2M gC3M gEM.

[0123] FIG. 17—Expected labeling of proteinogenic amino acids upon feeding with 13C-formate/12C-CO2or 12C-formate/13C-CO2 and according to different metabolic scenarios.

[0124] FIG. 18—Number of colony forming units increases monotonically with OD600 for cells growing on methanol as sole carbon source.

[0125] FIG. 19—Addition of 100 mM sodium bicarbonate increases final OD600 on methanol, reaching 0.9 instead of 0.2 (FIG. 5C). Consumption of methanol is depicted by the bars: the grey bars correspond to methanol concentration in a test tube without cells (concentration decrease due to evaporation), while the blue bars represent the concentration of methanol in a test tube in which cells are growing on methanol.

[0126] The examples illustrate the invention.

EXAMPLE 1—RESULTS

The Reductive Glycine Pathway

[0127] Escherichia coli, as most other key biotechnological microorganisms, cannot naturally grow on C.sub.1 feedstocks. In this study, it was aimed to design and engineer a simple, linear synthetic pathway which could support E. coli growth on formate or methanol as sole carbon source. The inspiration came from the anaerobic reductive acetyl-CoA pathway (rAcCoAP).sup.23 which assimilates C.sub.1 compounds very efficiently. The reductive glycine pathway (rGlyP), as shown in FIG. 1, was designed to be the aerobic twin of the rAcCoAP.sup.24. Both are linear routes with limited overlap with central metabolism, minimizing the need for regulatory optimization. Both pathways start with the ligation of formate and tetrahydrofolate (THF), proceed via reduction into a C.sub.1-THF intermediate, which is then condensed, within an enzyme complex, with CO.sub.2 to generate a C.sub.2 compound (acetyl-CoA or glycine). The C.sub.2 compound is finally condensed with another C.sub.1 moiety and metabolized to generate pyruvate as biomass precursor. Importantly, both the rAcCoAP and the rGlyP are characterized by a ‘flat’ thermodynamic profile.sup.24,25, that is, both are mostly reversible such that the direction of the metabolic flux they carry is determined mainly by the concentrations of their substrates and products. This thermodynamic profile, while constraining the driving force of the pathway reactions.sup.26, indicates very high energetic efficiency, where no energetic input, e.g., in the form of ATP hydrolysis, is wasted. Indeed, both pathways are associated with a very low ATP cost: only 1-2 ATP molecules are invested in the metabolism of formate to pyruvate.sup.24. Yet, unlike the rAcCoAP, the key enzymatic components of which are highly oxygen sensitive, the rGlyP can operate under full aerobic conditions. Hence, the rGlyP represents the most efficient theoretical route—in terms of energy utilization, resources consumption, and biomass yield—to assimilate formate in the presence of oxygen.sup.24.

[0128] A recent study suggests that the complete rGlyP might be naturally operating in a phosphite-oxidizing microbe.sup.27. Moreover, the key enzymatic conversion of the rGlyP, catalyzed by the glycine cleavage system (GCS), was shown to be fully reversible in many organisms.sup.28-30. Previous studies demonstrated that the GCS can support glycine and serine biosynthesis from formate in an engineered E. coli strain at elevated CO.sub.2 concentration.sup.31-33. However, growth of the microorganism on formate (and CO.sub.2) as a sole carbon source has not yet been demonstrated and remains an open challenge.

Modular-Engineering Approach Establishes Grow on Formate

[0129] To facilitate the establishment of formatotrophic growth, the rGlyP was divided into four metabolic modules (FIG. 6): (i) a C.sub.1 Module (C.sub.iM), consisting of formate THF ligase, methenyl-THF cyclohydrolase, and methylene-THF dehydrogenase, all from Methylobacterium extorquens.sup.34, together converting formate into methylene-THF; (ii) a C.sub.2 Module (C.sub.2M), consisting of the endogenous enzymes of the GCS (GcvT, GcvH, and GcvP) which condenses methylene-THF with CO.sub.2 and ammonia to give glycine; (iii) a C.sub.3 Module (C.sub.3M), consisting of serine hydroxymethyltransferase (SHMT) and serine deaminase, together condensing glycine with another methylene-THF to generate serine and finally pyruvate; and (iv) an Energy Module (EM), which consists of formate dehydrogenase (FDH) from Pseudomonas sp. (strain 101).sup.35, generating reducing power and energy from this C.sub.1 feedstock.

[0130] The strategy was to establish the activities of the different modules in consecutive steps, integrating subsequent modules and selecting for their combined activity. It was started with an E. coli strain that is auxotrophic for serine, glycine, and C.sub.1 moieties—ΔserA Δkbl ΔltaE ΔaceA—where the first deletion abolishes native serine biosynthesis, the second and third abolish threonine cleavage to glycine, and the final deletion prevents the formation of glyoxylate that could potentially be aminated to glycine.sup.32. The combined activity of the C.sub.1M and the C.sub.2M, together with the native activity of SHMT, should enable the cell to metabolize formate into C.sub.1-THF, glycine, and serine, relieving these auxotrophies (FIG. 2A).

[0131] Into the serine auxotroph strain, the enzymes of the C.sub.1M and the C.sub.2M were introduced, either on plasmid or in the genome (FIG. 6). For genome integration of C.sub.1M, all relevant enzymes were combined into one operon, under the regulation of a strong constitutive promoter.sup.36, which was inserted into a genomic ‘safe spot’, SS9.sup.37. In the case of the C.sub.2M, the native promoter of the GCS was replaced with a strong constitutive one (FIG. 6), increasing transcript levels 20-50 fold (FIG. 7). As expected, growth with formate was observed upon overexpression of both modules (FIG. 2B) and was dependent upon high CO.sub.2 concentration (10% in the headspace) which thermodynamically and kinetically supports the reductive activity of the GCS. While genomic integration of the enzymes of the C.sub.1M (gC,M) did not improve growth compared to plasmid expression (pC.sub.1M), replacing plasmid borne expression of the enzymes of the C.sub.2M (pC.sub.2M) with genomic overexpression (gC.sub.2M) supported a higher growth rate (FIG. 2B).

[0132] Next, it was aimed to establish formate as the primary carbon source, which requires high expression of the enzymes of the C.sub.3M to convert glycine into the central metabolism intermediate pyruvate (FIG. 2C). To enable formate assimilation to biomass, an energy source is required, which at this stage was chosen to be acetate. The TCA cycle can fully oxidize acetate to generate reducing power and energy, while the deletion of isocitrate lyase (ΔaceA) abolishes the activity of the glyoxylate shunt, thus preventing the cell from using this molecule as a carbon source. Growth should thus be dependent on formate assimilation via the rGlyP for biomass generation and acetate oxidation for the production of reducing power and energy (FIG. 2C).

[0133] The enzymes of the C.sub.3M were either overexpressed on a plasmid (pC.sub.3M) or in the genome (gC.sub.3M) (FIG. 6); in the latter case, the native glyA and sdaA were deleted and a synthetic operon harboring both genes under the regulation of a strong constitutive promoter was introduced into another genomic ‘safe spot’, SS7.sup.37. Overexpression of the enzymes of the C.sub.3M, within a strain that genomically expresses the enzymes of the C.sub.1M and the C.sub.2M, resulted on growth on formate and acetate (at 10% CO.sub.2) (FIG. 2D). Genomic expression of C.sub.3M supported more robust growth compared to the C.sub.3M expressed from plasmid. To confirm that the expression level of C.sub.3M does not constrain the growth rate, a strain was tested in which the expression of glyA and sdaA is controlled by a stronger ribosome binding site (RBS-A instead of RBS-C.sup.36). It was found that this strain grows rather poorly (FIG. 8), indicating that higher expression of these genes is deleterious.

[0134] Finally, it was aimed to introduce the EM such that formate can serve as sole carbon and energy source (FIG. 2E). Overexpression of FDH on a plasmid (FIG. 6), in the strain carrying the genes of the C.sub.1M, C.sub.2M and C.sub.3M in the genome, enables growth on formate (FIG. 9). However, when FDH was introduced into yet another genomic ‘safe spot’, SS10.sup.37, it failed to establish growth (FIG. 9), suggesting that the expression level of FDH was too low. Therefore, a strain was tested in which the genomic expression of FDH was controlled by a stronger ribosome binding site (RBS-A instead of RBS-C.sup.36, FIG. 6). This strain, carrying no plasmid, was able to grow on formate as a sole carbon and energy source (FIG. 2F and FIG. 9). Growth on formate was also observed in a test-tube and confirmed by recording monotonically increasing colony-forming units with increased OD (FIG. 10). This is the first case in which growth on formate was made possible in a microorganism that cannot assimilate C.sub.1 compounds natively.

Short-Term Evolution Improves Growth on Formate

[0135] To improve growth on formate it was decided to conduct a short term evolution experiment in fed batch mode. The engineered strain was cultivated in test tubes, where formate was added every 3-6 days, increasing the concentration in the medium by 30 mM (red arrows in FIG. 3A). Once cell turbidity reached an OD.sub.600 of 0.4, the cells were diluted to OD.sub.600 of 0.03-0.05 and started a new cycle of cultivation (FIG. 3A shows six typical cycles).

[0136] Within 13 cultivation cycles (≤40 generations), growth rate on formate was substantially improved (FIG. 3A), with the doubling time dropping from 65-80 h in the first two cycles to less than 10 h in the last cycle (FIG. 3B). Growth yield on formate also improved, from ≈1.5 gCDW/mol-formate in the first cycle to 2.3±0.2 gCDW/mol-formate in the last. This yield is similar to that of microorganisms growing autotrophically on formate via the Calvin cycle (3.2±1.1 gCDW/mol-formate.sup.38). The growth of the evolved bacterium on formate was directly coupled to a decrease in the concentration of the feedstock in the medium (FIG. 3C). Furthermore, as formatotrophy consumes protons (net oxidation and net assimilation both consume formic acid rather than formate), a direct correlation was observed between cell density and the pH of the medium (FIG. 11).

[0137] To better characterize growth on formate, growth experiments were conducted in 96-well plates, automatically measuring OD.sub.600 every ˜10 minutes. It was found that maximal cell density increased monotonically with increasing formate concentration from 10 mM to 150 mM (FIG. 3D). Similarly, the doubling time decreased monotonically with increasing formate concentration: from 17 hours with 10 mM formate to less than 8 hours at formate concentrations higher than 100 mM (FIG. 3D). The cellular toxicity of formate, which is attributed to inhibition of cytochrome c oxidase.sup.39 and dissipation of the proton motive force.sup.40, probably explains the increased lag time at formate concentrations of 109 mM and 153 mM, and the failure to grow at higher concentrations.

[0138] Adaptive laboratory evolution usually requires hundreds of generation to improve the fitness of E. coli in a substantial way.sup.41-43. The strain required less than 40 generations, presumably as the growth of the parent strain was so poor that a small number of mutations were sufficient to drastically improve fitness. To check whether this is indeed the case, multiple colonies of the evolved strain were isolated and their genomes were sequenced. Two mutations were found which occurred in all sequenced colonies (FIG. 12). The first was a single base-pair substitution in the 5′-UTR of the newly introduced FDH gene, which increased the level of transcript 2.5-fold (FIG. 13) and resulted in a 7.4-fold increase in formate oxidation activity in cell extract assays (FIG. 14). The second mutation was a single base-pair substitution in the promoter region of pntAB, which encodes for the membrane-bound transhydrogenase. This mutation increased transcript level by more than 13-fold (FIG. 13). The beneficial effect of these two mutations is to be expected, as the first increases energy supply to the cell from formate and the second increases the availability of NAPDH, a key cofactor for the activity of the rGlyP (consumed by methylene-THF dehydrogenase), the supply of which could limit pathway activity.

[0139] To confirm that the two mutations suffice to support the improved growth on formate, Multiplex Automated Genomic Engineering (MAGE.sup.44) was used to introduce these mutations into a non-evolved strain. It was found that while the parent strain could hardly grow in 96-well plates, the strain in which the two mutations were present displayed a growth profile almost identical to that of the evolved strain (FIG. 15). It was therefore concluded that overexpression of FDH and PntAB were sufficient to enable the observed improved growth on formate. By further optimizing cultivation conditions, it was found that addition of 100 mM sodium bicarbonate to the medium enabled the evolved strain, as well as the reconstructed strain, to grow at higher formate concentrations, tolerating even 300 mM (FIG. 16).

Carbon Labeling Confirms Pathway Activity and Shed Light on Cellular Fluxes

[0140] To confirm that growth on formate indeed proceeds via the rGlyP, carbon labeling experiments were performed. The cultures were fed with .sup.13C-formate/.sup.12CO.sub.2, .sup.12C-formate/.sup.13CO.sub.2, and .sup.13C-formate/.sup.13CO.sub.2, and measured the labeling pattern of proteinogenic amino-acids using liquid chromatography-mass spectrometry. The focus was on 7 amino-acids—glycine, serine, alanine, valine, proline, threonine, and histidine—which either directly relate to the activity of the rGlyP or originate from different parts of central metabolism, thus providing an indication of key metabolic fluxes.

[0141] As shown in FIG. 4, the amino acid labeling confirms the activity of the rGlyP. Specifically, feeding .sup.13C-formate/.sup.12CO.sub.2 resulted in single labeled glycine and double labeled serine and pyruvate (as indicated by the labeling of alanine). As valine—derived from two pyruvate molecules, one of which loses its carboxylic acid carbon—is mostly quadruple labeled, it was deduce that pyruvate is labeled in its two non-carboxylic carbons, as predicted for growth via the rGlyP (FIG. 17). Conversely, feeding .sup.12C-formate/.sup.13CO.sub.2 resulted, as expected, in single labeled glycine, serine and pyruvate. As valine is also single labeled, it was deduced that pyruvate is labeled in its carboxylic carbon, again confirming the activity of the rGlyP (FIG. 17). Upon feeding .sup.13C-formate/.sup.13CO.sub.2, all amino-acids were nearly-completely labeled, where the overall fraction of labeled carbon (marked above the bars in FIG. 4 in italics) is 97-98%, as expected by feeding with 99% .sup.13C-labeled formate and 99% .sup.13C-labeled CO.sub.2.

[0142] The labeling of threonine (derived from oxaloacetate) and proline (derived from 2-ketoglutarate) sheds light on the flux via the anaplerotic reactions and the TCA cycle. Specifically, if cyclic flux via the TCA cycle would predominate over anaplerotic flux, threonine and proline would be expected to be almost fully labeled upon feeding with .sup.13C-formate and almost fully unlabeled when feeding with .sup.13CO.sub.2 (FIG. 17). Conversely, if anaplerotic flux and non-cyclic flux would predominate over the cyclic flux, then threonine would be expected to be mostly double labeled on either .sup.13C-formate or .sup.13CO.sub.2 and proline would be expected to be mostly quadruple labeled on .sup.13C-formate and single labeled on .sup.13CO.sub.2 (FIG. 17). The results shown in FIG. 4 are thus consistent with high anaplerotic flux and low cyclic flux. This indicates that the cell obtains sufficient reducing power and energy from formate oxidation via FDH, and hence does not need to wastefully oxidize the assimilated carbons within pyruvate and acetate (i.e., investing cellular resources for C.sub.1 assimilation, only to completely oxidize the assimilated product).

Engineered Growth of E.coli on Methanol

[0143] Next, it was aimed to use the rGlyP for methanol assimilation. A single enzyme, methanol dehydrogenase (MDH), can convert methanol to formaldehyde, which can be oxidized to formate by the endogenous glutathione system.sup.45 (FIG. 5A). The expression of MDH can thus be regarded as the introduction of another module—Methanol Module (MM)—that serves to metabolize methanol to formate, while providing the cells with reducing power (FIG. 5B). NAD-dependent MDH from several organisms was tested: Bacillus stearothermophilus (BsMDH) .sup.19, Corynebacterium glutamicum (CgMDH) .sup.46, and Cupriavidus necator N-1 (CnMDH).sup.47, as well as two MDHs from Bacillus methanolicus (BmMDH2 and BmMDH3).sup.10,48 and an improved variant (BmMDH2*, carrying Q5L A363L modifications).sup.48 These MDH variants were expressed on plasmids in three genetic backgrounds: the parent strain (gC.sub.1M gC.sub.2M gC.sub.3M gEM), the evolved strain, and the parent strain to which the mutation within the promoter of the pntAB (FIG. 12) was introduced via MAGE. Overexpression of BsMDH supported growth on 600 mM methanol, which was most efficient in the latter strain (FIG. 5C) and somewhat poorer in the other strains (FIG. 5D). Growth was confirmed by observing monotonically increasing colony-forming units with increased OD (FIG. 18). The other MDH variants failed to support growth (FIG. 5D, final OD.sub.600 not higher than inoculation, as indicated by the brown dashed line).

[0144] To confirm that growth on methanol indeed depends on formaldehyde oxidation via the glutathione system, the endogenous genes encoding for S-(hydroxymethyl)glutathione dehydrogenase (ΔfrmA) were deleted in the above strains. The deletion was found to completely abolish growth on methanol (FIG. 5D), confirming the essentiality of the glutathione system to the observed growth. Moreover, overexpression of NAD-dependent formaldehyde dehydrogenase from Pseudomonas putida (PpFADH; SEQ ID NOs: 49 and 50), as demonstrated in a previous study .sup.12, or from Pseudomonas aeruginosa (PaFADH.sup.49; SEQ ID NOs: 51 and 52) did not improve growth on methanol (FIG. 5D), indicating that the endogenous glutathione system is sufficiently fast and that the rate limiting step lies in methanol oxidation.

[0145] To confirm that growth on methanol indeed proceed via the rGlyP, a carbon labeling experiment was performed. The cultures were fed with .sup.13C-methanol/.sup.12CO.sub.2 and the labeling pattern of the proteinogenic amino-acids described above was measured. The measured labeling pattern (FIG. 5E) was essentially identical to that observed with .sup.13C-formate/.sup.12CO.sub.2 (FIG. 4), confirming that growth on methanol takes place via the synthetic route.

[0146] Notably, the growth rate on methanol was considerably lower than that on formate—doubling time of 54±5.5 h. This can be attributed to the slow rate of methanol oxidation. The observed biomass yield was 4.2±0.17 gCDW / mole methanol, considerably lower than that of microorganisms naturally growing on methanol (7.2±1.2 gCDW/mol-methanol via the Calvin cycle, 12±1.6 gCDW/mol-methanol via the serine cycle, and 15.6±2.7 gCDW/mol-methanol via the RuMP cycle.sup.38). It is speculated that the low yield is also related to the slow rate of methanol oxidation: a low growth rate increases the proportional consumption of energy for cell maintenance, thus lowering biomass yield. Addition of 100 mM sodium bicarbonate significantly increased the final OD.sub.600, but the growth parameters did not improve: doubling time of 55±1 h and biomass yield of 4.2±0.1 gCDW/mol-methanol (FIG. 19, also showing methanol consumption during growth).

Conclusions

[0147] This study provides the first demonstration of synthetic formatotrophy and methylotrophy. It is shown that rational design alone can suffice to achieve such a goal, but that short term evolution can provide useful fine tuning to improve growth characteristics. Further improvement of growth on formate and methanol can be achieved via long term evolution or via the introduction of metabolic routes that bypass limiting reactions. For example, replacing NAD-dependent MDH with methanol oxidase might reduce biomass yield (as this enzyme dissipates reducing power) but could support a much higher growth rate, as it replaces a thermodynamically- and kinetically-limited reaction with a favorable and fast one. The C.sub.1 assimilating strains can be further engineered for the production of value-added chemicals. Especially interesting are chemicals that can be derived directly from the rGlyP intermediates or product, and can thus be produced with high yield and productivity. For example, lactate and isobutanol, both of which are derived from pyruvate, should be produced with high yield. Similarly, cysteine, which is derived from serine, a key pathway intermediate, might be an ideal product. Coupling the abiotic synthesis of formate and methanol with their microbial conversion to chemicals of interest will enable an integrated process for the valorization of CO.sub.2 into renewable commodities.

EXAMPLE 2—MATERIAL AND METHODS

Chemicals and Reagents

[0148] Primers were synthesized by Integrated DNA Technologies (IDT, Leuven, Belgium). PCR reactions were carried out either using Phusion High-Fidelity DNA Polymerase or Dream Taq. Restrictions and ligations were performed using FastDigest enzymes and T4 DNA ligase, respectively, all purchased from Thermo Fisher Scientific (Dreieich, Germany). Glycine, sodium formate, sodium formate-.sup.13C, methanol-.sup.13C were ordered from Sigma-Aldrich (Steinheim, Germany). .sup.13CO.sub.2 was ordered from Cambridge Isotope Laboratories, Inc. (Andover, Mass., USA).

Bacterial Strains

[0149] Wild type Escherichia coli strain MG1655 (F.sup.−λ.sup.−ilvG.sup.−rfb-50 rph-1) was used as the host for all genetic modifications. E. coli strain DH5α (F.sup.−, λ.sup.−,ϕ80/lacZΔM15, Δ(lacZYA-argF)U169, deoR, recA1, endA1, hsdR17(rK.sup.−mK.sup.+), phoA, supE44, thi-1, gyrA96, relA1) and E. coli strain ST18 (pro thi hsdR.sup.+ Tp.sup.r Sm.sup.r−; chromosome::RP4-2 Tc::Mu-Kan::Tn7λpir ΔhemA).sup.50 were used for cloning and conjugation procedures, respectively.

Genome Engineering

[0150] Gene knockouts were introduced in MG1655 by P1 phage transduction.sup.51. Single gene knockout mutants from the National BioResource Project (NIG, Japan).sup.52 were used as donors of specific mutations. For the recycling of selection marker (as the multiple gene deletions and integrations were required) all the antibiotic cassettes integrated into genome were flanked by FRT (Flippase Recongnition Target) sites. Cells were transformed with a flippase recombinase helper plasmid (FLPe, replicating at 30° C., Gene Bridges), which carries a gene encoding FLP which recombines at the FRT sites and removes the antibiotic cassette. Elevated temperature (37° C.) was subsequently used to cure the cell from the FLPe plasmid.

[0151] Exchange of E. coli native promoter with a synthetic one was performed by using PCR-mediated λ-Red recombination method. The synthetic promoter fused with FRT-flanked kanamycin resistance gene was cloned into the pZ vector and the DNA fragment was obtained by PCR amplification with primers containing 50 base pair homology for recombination. Recombinant E. coli MG1655 harboring λ-Red recombinase (pRed/ET, Gene Bridges) was cultivated at 30° C., and the expression of λ-Red recombinase was induced by the addition of 10 mM L-arabinose. Electro-competent cells were prepared by washing three times with ddH.sub.2O. The PCR product was introduced into E. coil expressing the λ-Red recombinase via electroporation. Mutants with exchanged promoter occurred via homologous recombination, selected on the LB agar plate containing 50 μg ml.sup.−1 kanamycin, and subsequently screened by colony PCR.

[0152] To enable genomic overexpression from a synthetic operon, conjugation based genetic recombination methods was adapted as previously described.sup.36. The synthetic operons were digested with Bcul and Notl, and ligated by T4 ligase into previously digested with the same enzyme pDM4 (with oriR6K) genome integration vector. This vector has two 600 bp homology region compatible with target spot, chloramphenicol resistance gene (camR), a levansucrase gene (sacB), and the conjugation gene traJl for the transfer of the plasmid. The resulting ligation products were used to transform chemically competent E. coli ST18 strains. Positive clones growing on chloramphenicol medium supplemented with 5-aminolevulinic acid (50 mg mi.sup.−1) were identified by colony PCR, and the confirmed recombinant ST18 strain was used as donor strain for the conjugation. Chloramphenicol resisting recipient E. coil strains were screened as positive strains for the first round of recombination. Subsequently, sucrose counter selection and kanamycin resistance tests were carried out to isolate recombinant E. coli strains with the correct synthetic operon integration into chromosome. All constructs were verified via PCR and sequencing.

[0153] Introducing point mutations on genome—to establish the mutation shown in FIG. 12—was achieved by using multiplex automated genome engineering (MAGE).sup.44,53. A single colony of desired strain(s) transformed with pORTMAGE.sup.53 (Addgene catalog no. 72680) was incubated in LB medium supplemented with 100 mg I.sup.−1 of ampicillin at 30° C. in a shacking incubator. To start the MAGE cycle, overnight cultures were diluted by 100 times in the same medium and cultivated to an optical density of 0.4-0.5 at 600 nm. 1 ml of each culture was transferred to sterile microcentrifuge tubes, and then transferred to 42° C. thermomixer (Thermomixer C, Eppendorf) to express λ-Red genes by heat shock for 15 min at 1000 rpm. After induction, cells were quickly chilled on ice for at least 15 min, and then made electrocompetent by washing three times with ice-cold ddH.sub.2O. 40 ul of electrocompetent cell was mixed with 2 ul of 50 uM of oligomer stock solution and the final volume of the suspension was adjusted to 50 ul. The oligomers used for MAGE were: 5″-T*T*T TTG GCG CTA GAT CAC AGG CAT AAT TTT CAG TAC GTT ATA GGG tGT TTG TTA CTA ATT TAT TTT AAC GGA GTA ACA TTT AGC TCG T*A*C -3″ (pntAB_MAGE; SEQ ID NO: 53), 5′-T*A*A AGT TAA ACA AAA TTA TTT CTA TTA ACT AGT GAA TTC GGT CAt TGC GTC CTG CGC ATA TTA TAT GTG AAT CAC AGT GAT ATG TCA A*G*T-3′ (fdh_MAGE; SEQ ID NO: 54) where the asterisk (*) indicates phosphorothiolated bond. Electroporation was done on Gene Pulser XCell (Bio-Rad) set to 1.8 kV, 25 μF capacitance, and 200 Ω resistance for 1 mm gap cuvette. Immediately after electroporation, 1 ml of LB was added to cuvette and the electroporation mixes in LB was transferred to sterile culture tubes and cultured with shaking at 30° C., 240 rpm for 1 hour to allow for recovery. After recovery, 2 ml of LB medium supplemented with ampicillin was added and then further incubated in the same condition. When the culture reached an OD.sub.600 of 0.4-0.5, cells were either subjected to additional MAGE cycle or analyzed for genotype via PCR and sequencing. 8 consecutive MAGE cycles were performed before analyzing the genotype to identify strains carrying the required mutations.

[0154] All strains used are shown in Table 1.

TABLE-US-00001 TABLE 1 Strains and plasmids used in this study Strain/Plasmid Description/Genotype Source Strains MG1655 F.sup.− λ.sup.− ilvG.sup.− rfb-50 rph- 1 DH5α F.sup.− λ.sup.− Φ80lacZΔM15 Δ(lacZYA-argF)U169 deoR recA1 2 endA1 hsdR17(rK.sup.− mK.sup.+) phoA supE44 thi-1 gyrA96 relA1 ST18 pro thi hsdR.sup.+ Tp.sup.r Sm.sup.r; chromosome::RP4-2 Tc::Mu- 3 Kan::Tn7/λpir ΔhemA SerAux MG1655, ΔserA ΔltaE Δkbl ΔaceA 4 gC.sub.1M SerAux, ss9-P.sub.STRONG-RBS.sub.C-ftfL-RBS.sub.C-fch-RBS.sub.C-mtdA This study gC.sub.2M SerAux, P.sub.STRONG-RBS.sub.C-gcvT-RBS.sub.NATIVE-gcvH-RBS.sub.NATIVE- This gcvP study gCM gC.sub.2M gC.sub.1M, P.sub.STRONG-RBS.sub.C-gcvT-RBS.sub.NATIVE-gcvH-RBS.sub.NATIVE- This gcvP study gC.sub.1M gC.sub.2M gC.sub.3M gC.sub.1M gC.sub.2M, ss7-P.sub.STRONG-RBS.sub.C-glyA-RBS.sub.C-sdaA ΔsdaA This ΔglyA study gC.sub.1M gC.sub.2M gC.sub.3M’ gC.sub.1M gC.sub.2M, ss7-P.sub.STRONG-RBS.sub.A-glyA-RBS.sub.A-sdaA ΔsdaA This ΔglyA study gC.sub.1M gC.sub.2M gC.sub.3M gC.sub.1M gC.sub.2M gC.sub.3M, ss10-P.sub.STRONG-RBS.sub.A-fdh This gEM (K4) study gC.sub.1M gC.sub.2M gC.sub.3M gC.sub.1M gC.sub.2M gC.sub.3M, ss10-P.sub.STRONG-RBS.sub.C-fdh This gEM' study K4 g-PntAB* K4 strain with a point mutation in promoter region of pntAB This study K4 g-FDH* g- K4 strain with a point mutation in both promoter region of This PntAB* pntAB and 5’UTR region of ss10-P.sub.STRONG-RBS.sub.A-fdh study K4e Evolved K4 strain after short term evolution This study Plasmids pDM4 Conjugation plasmid with oriR6K origin, sacB, traJI and 5 chloramphenicol/kanamycin resistance pZASS Overexpression plasmid with p15A origin, streptomycin 5 resistance, constitutive strong strength promoter (P.sub.STRONG) pZASM Overexpression plasmid with p15A origin, streptomycin 5 resistance, constitutive medium strength promoter (P.sub.MEDIUM) pZATM Overexpression plasmid with p15A origin, tetracycline 5 resistance, constitutive medium strength promoter (P.sub.MEDIUM) pZSSM Overexpression plasmid with pSC101 origin, streptomycin 5 resistance, constitutive medium strength promoter (P.sub.MEDIUM) pDM4:SS9-C.sub.1M pDM4 backbone with 600 bp up/down homology to safe This spot 9 .sup.6 for the genome integration of P.sub.STRONG-RBS.sub.C-ftfL- study RBS.sub.C-fch-RBS.sub.C-mtdA pDM4:SS7-C.sub.3M pDM4 backbone with 600 bp up/down homology to safe This spot 7 .sup.6 for the genome integration of P.sub.STRONG-RBS.sub.C-glyA- study RBS.sub.C-sdaA pDM4:SS10-EM pDM4 backbone with 600 bp up/down homology to safe This spot 10 .sup.6 for the genome integration of P.sub.STRONG-RBS.sub.A-fdh study pC.sub.1M pZSSM backbone for overexpression of RBS.sub.C-ftfL-RBS.sub.C- 4 fch-RBSc-mtdA from Methylobacterium extorquens pC.sub.2M pZATM backbone for overexpression of RBS.sub.C-gcvT-RBS.sub.C- 4 gcvH-RBS.sub.C-gcvP from E. coli pC.sub.3M pZASS backbone for overexpression of RBS.sub.C-glyA-RBS.sub.C- This sdaA from E. coli study ASS-glyA-sdaA pZASS backbone for overexpression of RBS.sub.C-glyA-RBS.sub.C- This sdaA from E. coli study ASM-glyA-sdaA pZASS backbone for overexpression of RBS.sub.C-glyA-RBS.sub.C- This sdaA from E. coli study ASS-sdaA pZASS backbone for overexpression of RBS.sub.C-sdaA from This E. coli study ASM-sdaA pZASM backbone for overexpression of RBS.sub.C-sdaA from This E. coli study ASS-fdh pZASS backbone for overexpression of RBS.sub.C-fdh from This Pseudomonas putida study ASS-bsMDH pZASS backbone for overexpression of methanol This dehydrogenase from Bacillus stearothermophilus study (UnitProt, P42327) ASS-cgMDH pZASS backbone for overexpression of methanol This dehydrogenase from Corynebacterium glutamicum study (UnitProt, A4QHJ5) ASS-cnMDH pZASS backbone for overexpression of methanol This dehydrogenase from Cupriavidus necator (UnitProt, study F8GNE5) ASS-bmMDH3 pZASS backbone for overexpression of methanol This dehydrogenase from Bacillus methanolicus (Unitprot, study I3E2P9) ASS-bmMDH2 pZASS backbone for overexpression of methanol This dehydrogenase from Bacillus methanolicus (Unitprot, study I3E949) ASS-bmMDH2* pZASS backbone for overexpression of engineered This methanol dehydrogenase (Q5L A363L) from Bacillus study methanolicus (Unitprot, I3E949) ASS- pZASS backbone for overexpression of RBSc-bsMDH- This bsMDH/paFADH RBSc-paFADH, a formaldehyde dehydrogenase from study Pseudomonas aeruginosa ASS- pZASS backbone for overexpression of RBSc-bsMDH- This bsMDH/ppFADH RBSc-ppFADH, a formaldehyde dehydrogenase from study Pseudomonas putida

REFERENCES IN TABLE Table 1

[0155] 1 Blattner, F. R. et al. The complete genome sequence of Escherichia coli K-12. Science 277, 1453-1462 (1997).

[0156] 2 Meselson, M. & Yuan, R. DNA restriction enzyme from E. coli. Nature 217, 1110-1114 (1968).

[0157] 3 Thoma, S. & Schobert, M. An improved Escherichia coli donor strain for diparental mating. FEMS Microbiol Lett 294, 127-132, doi:10.1111/.1574-6968.2009.01556.x (2009).

[0158] 4 Yishai, O., Bouzon, M., Doring, V. & Bar-Even, A. In Vivo Assimilation of One-Carbon via a Synthetic Reductive Glycine Pathway in Escherichia coli. ACS synthetic biology, doi:10.1021/acssynbio.8b00131 (2018).

[0159] 5 Wenk, S., Yishai, O., Lindner, S. N. & Bar-Even, A. An Engineering Approach for Rewiring Microbial Metabolism. Methods Enzymol 608, 329-367, doi:10.1016/bs.mie.2018.04.026 (2018).

[0160] 5 Bassalo, M. C. et al. Rapid and Efficient One-Step Metabolic Pathway Integration in E. coli. ACS synthetic biology 5, 561-568, doi:10.1021/acssynbio.5b00187 (2016).

Synthetic-Operon Construction

[0161] Protein sequences of formate-tetrahydrofolate ligase (ftfL, UniProt: Q83WS0), 5,10-methenyl-tetrahydrofolate cyclohydrolase (fchA, UniProt: Q49135), and 5,10-methylene-tetrahydrofolate dehydrogenase (mtdA, UniProt: P55818) were taken from Methylobacterium extorquens AM1. Formate dehydrogenase (fdh, UniProt: P33160) was taken from Pseudomonas sp. Formaldehyde dehydrogenase were obtained from Pseudomonas aeruginosa (fdhA, UnitProt: Q9HTE3) and Pseudomonas putida (fdhA, UnitProt: P46154). Methanol dehydrogenases were prepared from Bacillus stearothermophilus (adh, UniProt: P42327), Corynebacterium glutamicum (cgR_2695, UniProt: A4QHJ5), Cupriavidus necator (mdh2, UniProt: F8GNE5), and Bacillus methanolicus (UnitProt: I3E2P9 and I3E949, as well as en engineered MDH, as reported in.sup.48). These genes were codon optimized for E. coli K-12 and synthesized (Baseclear, Netherlands). All gene sequences are listed in sequence protocol of the application.

[0162] Genes native to E. coli—that is, serine hydroxymethyltransferase (glyA) and serine deaminase (sdaA)—were prepared via PCR-amplification from E. coli MG1655 genome. Genes were integrated into a high copy number cloning vector pNiv to construct synthetic operons using the method described previously.sup.36,54 Plasmid-based gene overexpression was achieved by cloning the desired synthetic operon into the pZ vector (15A origin of replication, streptomycin marker.sup.36) digested with EcoRI and Pstl utilizing T4 DNA ligase. All molecular biology techniques were performed with standard methods.sup.55 or following manufacturer protocol.

[0163] Promoters and ribosome binding sites were used as described previously.sup.36,54,56. Briefly, either a medium strength constitutive promoter (‘PGl-10’.sup.56) or a strong constitutive promoter (‘PGl-20’.sup.56) was used, as indicated in the text and in FIG. 6. Either medium strength ribosome binding site (RBS.sub.c.sup.54) or a strong ribosome binding site (RBS.sub.A.sup.54) was further used, as indicated in the text and in FIG. 6.

[0164] All plasmid used are shown in the above Table 1.

Growth Medium and Conditions

[0165] Luria Bertani medium (1% NaCl, 0.5% yeast extract, and 1% tryptone) was used for strain propagation. Further cultivation was done in M9 minimal media (50 mM Na.sub.2HPO.sub.4, 20 mM KH.sub.2PO.sub.4, 1 mM NaCl, 20 mM NH.sub.4Cl, 2 mM MgSO.sub.4, and 100 μM CaCl.sub.2), with trace elements (134 μM EDTA, 13 μM FeCl.sub.3.6H.sub.2O, 6.2 μM ZnCl.sub.2, 0.76 μM CuCl.sub.2.2H.sub.2O, 0.42 μM CoCl.sub.2.2H.sub.2O, 1.62 μM H.sub.3BO.sub.3, 0.081 μM MnCl.sub.2.4H.sub.2O). For the cell growth test, overnight cultures in LB medium were used to inoculate a pre-culture at an optical density (600 nm, OD.sub.600) of 0.02 in 4 ml fresh M9 medium containing 10 mM glucose, 1 mM glycine and 30 mM formate in 10 ml glass test tube. Cell were then cultivated at 37° C. and shaking of 240 rpm. Cell cultures were harvested by centrifugation (18,407×g, 3 min, 4° C.) and washed twice with fresh M9 medium and used to inoculate the main culture, conducted aerobically either in 10 ml glass tube or Nunc 96-well microplates (Thermo Fisher Scientific) with appropriate carbon sources according to strain and specific experiment: 10 mM glucose, 20 mM acetate, 30 mM formate, 600 mM methanol, and/or 10% CO.sub.2 (90% air). In the microplates cultivation, each well containing 150 pl culture covered with 50 μl mineral oil (Sigma-Aldrich) to avoid evaporation (note that small gaseous molecules such as O.sub.2 and CO.sub.2 can freely diffuse via this oil coverage). Growth experiments were conducted (either 100% air or 90% ai/10% CO.sub.2) in a BioTek Epoch 2 plate reader (BioTek Instrument, USA) at 37° C. Growth (OD.sub.600) was measured after a kinetic cycle of 12 shaking steps, which alternated between linear and orbital (1 mm amplitude), and were each 60 s long. OD values measured in the plate reader were calibrated to represent OD values in standard cuvettes according to ODcuvette=ODplate/0.23. Glass tube culture was carried out in 4 ml of working volume, at 37° C. and shaking of 240 rpm. Volume loss due to evaporation was compensated by adding the appropriate amount of sterile double distilled water (ddH.sub.2O) to culture tube every two days. All growth experiments were performed in triplicate, and the growth curves shown represent the average of these triplicates.

.SUP.13.C Labeling of Proteinogenic Amino Acids

[0166] For stationary isotope tracing of proteinogenic amino acids, cells were cultured in 4 ml of M9 media supplemented with either labeled or unlabeled carbon sources, that is, .sup.13C-formate, .sup.13C-methanol and/or .sup.13CO.sub.2, under conditions as described above. A 6 L vacuum desiccator (Lab Companion, South Korea) was used for cultures grown in .sup.13CO.sub.2, where the original gas was expelled by using vacuum pump followed by refilling with 90% air and 10% .sup.13CO.sub.2. The cell was harvested by centrifugation for 3 min at 18,407×g when the stationary growth phase was reached. Biomass was hydrolyzed by incubation with 1 ml of 6 N hydrochloric acid for a duration of 24 h in 95° C. Samples were dried via heating at 95° C. and re-dissolved in 1 ml of ddH.sub.2O. Hydrolyzed amino acids were separated using ultra performance liquid chromatography (Acquity, Waters, Milford, Mass., USA) using a C18-reversed-phase column (Waters) as previously described.sup.57. Mass spectra were acquired using an Exactive mass spectrometer (Thermo Fisher). Data analysis was performed using Xcalibur (Thermo Fisher). Prior to analysis, amino-acid standards (Sigma-Aldrich) were analyzed under the same conditions in order to determine typical retention times.

Dry Weight Analysis

[0167] To determine dry cell weight of E. coil grown formate or methanol, pre-cultures prepared as described above were inoculated to at a final OD.sub.600 of 0.01 into fresh M9 medium containing either formate (30 mM) or methanol (600 mM) in 125 ml pyrex Erlenmeyer flask and grown at 37° C. with agitation at 240 rpm. Up to 50 ml of cell culture, growing in shake-flasks, were harvested by centrifugation (3,220×g, 20 min). To remove residual medium compounds cells were washed be three cycles of centrifugation (7,000×g, 5 min) and resuspension in 2 ml ddH.sub.2O. Cell-solutions were transferred to pre-weighted and pre-dried aluminum dish, dried at 90° C. for 16 h, and weight of the dried cells in the dish was determined and subtracted by the weight of the empty dish.

[0168] CDW of E. coil strains was measured during exponential growth phase (OD.sub.600 of 0.3-0.4) in the presence of 10% CO.sub.2 on 30 mM formate (at OD.sub.600 of 0.2, 0.37, and 0.41) and on 600 mM methanol (at OD.sub.600 of 0.21, 0.22, and 0.24). As a control, CDW of E. coli strain growing either on formate or methanol was determined during exponential growth phase in the presence of 10% CO.sub.2 and 30 mM formate and either 10 mM glucose (at OD.sub.600 of 1.26), 20 mM pyruvate (at OD.sub.600 of 0.78), or 20 mM succinate (at OD600 of 0.37). To determine CDW of E. coli WT, cells were grown in the presence of 10% CO.sub.2 on 10 mM glucose and CDW was determined during exponential growth phase (at OD.sub.600 of 0.78).

Enzymes and Chemical Assays

[0169] Absorbance changes for all assays were monitored in a BioTek Epoch 2 plate reader. Working at the measurement linear range was confirmed in all assays. Results represent averages of at least three cell preparations. To determine the activity of formate dehydrogenase, 1.5 ml of OD.sub.600 1.0 cell culture grown in M9 minimal medium and supplemented with glucose and formate from glass test tubes were washed twice with 9 gl.sup.−1 sodium chloride. Cells were lysed by adding CelLytic Reagent (Sigma) and allowed to sit for 20 min at the room temperature. After cell disruption, cellular debris was removed by centrifugation (18,407×g, 4° C., 10 min) and the supernatant used for crude assays without further purification. Formate dehydrogenase assay performed in the presence of 10 mM 2-mercaptoenthanol, 100 mM sodium formate, 200 mM sodium phosphate buffer pH 7.0, and 2 mM NAD.sup.+ in a total volume of 200 μl at 37° C..sup.58. The increase in NADH concentration resulting from formate oxidation was monitored at 340 nm. Protein concentration was measured using the Bradford Reagent (Sigma) with bovine serum albumin as a standard. Formate and methanol in the culture were quantified by a colorimetric assay using formate assay kit (Sigma-Aldrich) and methanol assay kit (BioVision) respectively. All samples were diluted to ensure the reading are within the standard curve range according to the manufacturer's instructions.

Quantitative Polymerase Chain Reaction

[0170] Total RNA was extracted from 1 ml of overnight culture at an OD.sub.600 0.5 using the RNeasy Mini Kit (Qiagen, Hilden, Germany), and following the protocol of the supplier. All RNA samples were treated with DNase I (Sigma-Aldrich, St. Louis, Mo., US) to remove any residual DNA. First-strand cDNA was synthesized using a qScript cDNA Synthesis kit following the manufacturer instructions (Quanta Biosciences, Gaithersburg, Md., US), and 1 μg of total RNA was used as a template in 20 μl reaction volume. Quantitative reverse-transcription-polymerase chain reactions (qRT-PCR) were made using a Maxima™ SYBR Green qPCR Master Mix (ThermoFisher Scientific, Darmstadt, Germany) supplemented with 5 μM primers and 5 μl cDNA template, which was diluted up to 200 μl after synthesis. The primers used for QPCR were: GCC AAT CTG CAA CAG TGC TC-3′ (pntA_forward, SEQ ID NO: 55), 5′-TTT TTG GCT GGA TGG CM GC-3′ (pntA_reverse, SEQ ID NO: 56), 5″- CGT GAC GM TAC CTG ATC GTT -3′ (fdh forward, SEQ ID NO: 57), 5″- GGT AGC GTT ACC TTT AGA GTA AGA GTG -3′ (fdh reverse, SEQ ID NO: 58). PCR was performed in 96-well optical reaction plates (ThermoFisher Scientific, Darmstadt, Germany) as follows: 10 min at 50° C., 5 min at 95° C., and 40 cycles of 10 s at 95 and 30 s at 60° C., and finally 1 min at 95° C. The specificity of the reactions, and the amplicon identities were verified by melting curve analysis. Reaction mixtures without cDNA were used as a negative control. Data were evaluated using the CT method.sup.59 and with correction for the PCR efficiency, which was determined based on the slope of standard curves. Normalization of gene expression levels were carried on by using the rrsA gene.sup.60, and eventually the fold-differences in the transcript levels and mean standard error were calculated as described elsewhere.sup.59.

Quantification of E. coil Colony Forming Units

[0171] Viable cell counts were determined by sampling E. coil cell cultures periodically. 10 μl of cell culture was diluted in 990 μl sterile M9 medium, and the diluted cell suspension was further diluted either by 100 times or 1000 times to obtain isolated colonies on agar plates. 100 μl of repeatedly diluted cell suspension was plated on LB agar plate and incubated at 37° C. for 24 h. All cell counts experiments were conducted at least five times per each OD value to obtain reliable cell counting numbers.

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