Bringing Carbon to Life via One-Carbon Metabolism

Abstract

One-carbon (C1) compounds found in greenhouse gases and industrial waste streams are underutilized carbon and energy sources. While various biological and chemical means exist for converting C1 substrates into multi-carbon products, major challenges of C1 conversion lie in creating net value. Here, we review metabolic strategies to utilize carbon across oxidation states. Complications arise in biochemical C1-utilization approaches because of the need for cellular energy currency adenosine triphosphate (ATP). ATP supports cell maintenance and proliferation and drives thermodynamically challenging reactions by coupling them with ATP hydrolysis. Powering metabolism through substrate cofeeding and energy transduction from light and electricity improves ATP availability, relieves metabolic bottlenecks, and upcycles carbon. We present a bioenergetic, engineering, and technoeconomic outlook for bringing elements to life.

Motivation and Challenges for Upcycling One-Carbon Compounds

One-carbon (C1) compounds are gaining traction as sustainable feedstocks to produce valuable chemicals. C1 compounds are often found in waste streams, and their conversion to value-added products offers environmentally beneficial alternatives to conventional chemical processes, which start from fossil resources and emit greenhouse gases. Methane and carbon dioxide (CO₂) are greenhouse gases which pose a global threat. Carbon monoxide (CO) is a poisonous gas found in syngas and partial combustion. Formate, formaldehyde, and methanol are C1 intermediates of methane, CO, and CO₂ conversion. Various technologies for utilizing C1 substrates exist today. Electrolysis uses electric potential to push redox reactions that are otherwise non-spontaneous. By applying electric potential, CO₂ can be converted to other C1 products [1, 2] and, with serial reduction, simple multi-carbon products such as acetate, ethanol and ethylene [3, 4]. Bioconversion, on the other hand, uses natural and synthetic pathways consisting of enzyme-catalyzed reaction steps to convert C1 feedstocks into biomass and a plethora of bioproducts.

Cellular metabolism encompasses biochemical reactions responsible for the synthesis of biomass and bioproducts as well as the generation of cellular energy currency adenosine triphosphate (ATP) and reducing power. Metabolic engineering endeavors rewire cellular metabolism to confer organisms with new and improved bioconversion capabilities. Many C1-utilizing organisms exist in nature, and their metabolism has been mapped [5–8]. Synthetic and systems biology tools have contributed to understanding and harnessing the full potential of metabolic pathways [9]. Different metabolic pathways are distinctively suitable for accommodating substrates and products with bioenergetic feasibility and biochemical proximity in terms of the number of reactions steps. Thus, for efficient and economically viable bioconversion, metabolic engineering is accompanied by a careful pairing of substrate and product [10].

Like any other system, operating metabolism costs energy. Thus, we asked three questions: Where does energy come from? How can energy be supplied without interruption? Does the output value exceed the input energy cost? Fortunately, cells can power metabolism using chemical, electrical, and light energy. Nutrient environments dictate how cells generate usable energy ATP and reducing equivalents from organic molecules. Metabolism works in synergy with abiological systems such as electrochemical cells and light-harvesting materials. Nonetheless, creating value while upcycling C1 compounds remains challenging. Here, we outline the pathways of one-carbon metabolism, diverse energy provision strategies, and biochemical and energetic considerations for efficient C1 conversion. Energy diversification and efficiency maximization principles will engender market-favored green technologies that outcompete conventional chemical processes.

Main Text

Physicochemical Considerations for C1 Compound Utilization

Efficient conversion of C1 compounds goes hand in hand with efficient energy transduction. Comparison of the oxidation states between C1 substrates and desired products indicates how many electrons need to be added or moved for the conversion. In electrolysis, an electric potential is applied to provide a driving force for electron transfer. In biology, enzymes interconvert and assimilate C1 substrates (Figure 1a). While oxidation number does not capture the thermodynamic, kinetic, and mass transport phenomena [2, 3, 11], it alludes to potential intermediates and side products. It also indicates whether a substrate can be an energy source for living organisms. For example, carbon in its most oxidized state, CO₂, can be building blocks for biomolecules, but energy needs to be supplied to cells for the assimilation of CO₂ into metabolism.

Figure 1. Energetics of different one-carbon substrates.

(a) Comparison between the oxidation numbers of one-carbon (C1) substrates and the average oxidation numbers of the carbons of metabolites and bioproducts. The fatty acid oxidation numbers range from 0 (acetic acid, a short-chain fatty acid) to ~2 (very-long-chain fatty acids). The amino acids oxidation number indicates the average of the carbons of the 20 canonical amino acids [94]. Enzymatic reactions that facilitate interconversion between C1 substrates include: MMO, methane monooxygenase; MDH, methanol dehydrogenase; MOX, methanol oxidase; FDH, formate dehydrogenase. (b) Comparison of the oxidation number of the carbon and the number of known major pathways for each substrate. (c) Comparison of the oxidation number of the carbon and number of ATP molecules required to produce convergent central carbon metabolites on a per-carbon basis for each C1 substrate. CH₄ represents methane; CH₃OH, methanol; CH₂O, formaldehyde; CO₂, carbon dioxide; HCOOH, formic acid; MMO, methane monooxygenase; MDH, methanol dehydrogenase; MOX, methanol oxidase; CODH, carbon monoxide dehydrogenase; FADH, formaldehyde dehydrogenase; FDH, formate dehydrogenase; RuMP, ribulose monophosphate cycle; EuMP, erythrulose monophosphate cycle; FORCE, formyl-CoA elongation reactions; HSC, homoserine cycle; WLP, Wood-Ljungdahl pathway; rGlyP, reductive glycine pathway; SerC, Serine Cycle; and CBB, Calvin-Benson-Bassham cycle.

C1 substrates can be assimilated via several metabolic pathways (Figure 1b) with varying ATP requirements (Figure 1c). The presence of C1 assimilation pathways does not immediately render them feasible alternatives to conventional chemical processes. Despite the Wood-Ljungdahl pathway (WLP) in acetogenic bacteria for converting CO₂, CO, and formate into acetate and downstream commodity chemicals, substantial bioenergetic constraint exists [12]. During gas fermentation, acetogens are ATP-limited due to diffusion and solubility limitations as well as a lack of substrate-level phosphorylation [13]. Thus, ATP generation is a key engineering objective for biochemical C1 upcycling.

Biochemical Considerations for Pairing C1 Compounds and Sensible Bioproducts

From a biological perspective, an ideal C1 substrate provides the carbon backbone for not only bioproducts but also biomass precursors (e.g., amino acids, nucleotides, and lipids). One-carbon metabolism condenses C1 substrates to two-carbon (C2) and/or three-carbon (C3) metabolites. Each C1 metabolic pathway has a unique cofactor (e.g., ATP and redox pairs) requirement and thermodynamics (Table 1) for assimilation into central carbon metabolism (Figure 2). From the resulting C2 and C3 metabolites, sensible bioproducts to synthesize are those that avoid as much oxidation and decarboxylation as possible.

Table 1:

C1 Metabolic pathway summary including substrates required and resulting products, examples of organisms in which the pathway occurs naturally, and Gibbs free energy of the overall reaction (ΔG’°).

Pathway	Substrates	Products	Relevant organisms	∆G’° (kJ/mol)	References
Wood-Ljungdahl Pathway (WLP)	4H2 + 2CO2	CH3COOH + 2H2O	Acetogens e.g., Moorella thermoacetica, Clostridium ljungdahlii, Acetobutylicum woodii, etc.	−95	[5]
Calvin Cycle (CBB) to Glucose	6CO2 + 12NADPH + 12H2O + 18ATP	C6H12O6 + 12NADP+ + 18ADP + 18Pi	Plants, cyanobacteria, algae, etc.	−358.8	[91]
Calvin Cycle (CBB)	3CO2 + 6NADPH + 9ATP + 5H2O	GAP + 6NADP + 9ADP + 8Pi	Plants, cyanobacteria, algae, etc.	−141.8	[92]
Reductive Glycine Pathway (rGlyP)	CHOOH + ATP + (THF) + 2NADH + NH3 + CO2	ADP + Pi + H2O + 2NAD+ + (THF) + C2H5NO2	Desulfovibrio desulfuricans, Saccharomyces cerevisiae, etc.	−17.2	[7, 29] eQuilibrator (equilibrator. weizmann.ac.il)
Serine Cycle	HCHO + CO2 + 2ATP + 2NADH + CoA	Acetyl-CoA + 2ADP + 2Pi + 2NAD+	Type II Methanotrophs e.g., Hyphomicrobium methylovorum GM2, Methylobacter whittenburyk, Methylocystis parvus, etc.	−123.5	[8, 40, 93] eQuilibrator (equilibrator. weizmann.ac.il)
Ribulose Monophosphate Cycle (RuMP)	3CH2O + ATP	DHAP + ADP	Type I Methanotrophs e.g., Bacilus subtilis, Methylobacillus flagellates, Mycobacterium gastri MN19, etc.	−89.9	MetaCyc (MetaCyc.org), eQuilibrator (equilibrator. weizmann.ac.il)

Figure 2. Assimilation of C1 compounds into central metabolism.

The oxidation states of C1 compounds and products are shown along metabolic reaction steps and pathways. CH₄ represents methane; CH₃OH, methanol; CH₂O, formaldehyde; CO₂, carbon dioxide; HCOOH, formic acid; CH₃COOH, acetic acid; C₂H₅NO₂, glycine; C₂H₄O₃, glycolic acid; MMO, methane monooxygenase; MDH, methanol dehydrogenase; MOX, methanol oxidase; CODH, carbon monoxide dehydrogenase; FADH, formaldehyde dehydrogenase; FDH, formate dehydrogenase; RuMP, ribulose monophosphate cycle; EuMP, erythrulose monophosphate cycle; FORCE, formyl-CoA elongation reactions; WLP, Wood-Ljungdahl pathway; rGlyP, reductive glycine pathway; and CBB, Calvin-Benson-Bassham cycle.

Photosynthetic organisms including plants and algae, the most familiar assimilators of CO₂, use the Calvin-Benson-Bassham (CBB) cycle to generate multi-carbon intermediates (Figure 3a) [14, 15]. The CBB cycle also enables utilization of CO in organisms that express carbon monoxide dehydrogenases, which may simultaneously provide electrons for respiration [16]. Interestingly, the genes for the CBB cycle are found in 6% of microbes whose genomes are sequenced [6], suggesting it could be engineered in non-photosynthetic organisms. Despite its relative prominence, the CBB cycle requires 3 ATP and 2 NADPH for reducing each CO₂ molecule. Ribulose bisphosphate carboxylase/oxygenase (RuBisCO) catalyzes the carboxylation step in the CBB cycle, but due to its slow turnover, it remains a bottleneck for the wider application of the pathway [6]. Introducing CBB genes to tractable model organisms may increase carbon yield by limiting net CO₂ loss. Heterologous expression of RuBisCO and phophoribulokinase (PRK) in E. coli in conjunction with its central carbon metabolism forms the CBB cycle, but regulatory and kinetic effects must also be considered for the CBB cycle to turn and fix CO₂ [17]. The expression of CBB proteins in E. coli can be negatively impacted by protein aggregates termed inclusion bodies (IB), which result from overexpressing foreign or mutated genes. IB formation is sensitive to environmental conditions; it has been shown that reducing the culture temperature from 37°C to 30°C results in uninterrupted CBB-cycle protein expression and increases CO₂ recycling to glyceraldehyde-3-phosphate (GAP) and subsequently lower glycolytic intermediates. Reducing the culture temperature of the CBB-capable E. coli strain led to a 2.3-fold increase in pyruvate production [18]. Using the C3 product of the CBB cycle to synthesize bioproducts that are derived from downstream C2 metabolites (e.g., acetyl-CoA and acetaldehyde) rather than C3 metabolites would lower carbon yield.

Figure 3. Major C1 assimilation pathways and potential products.

(a) The Calvin-Benson-Bassham Cycle (CBB) converts three CO₂ to glyceraldehyde 3-phosphate (GAP). (b) The Wood-Ljungdahl pathway (WLP) converts two CO₂ to acetyl-CoA. (c) The reductive glycine pathway (rGlyP) converts two CO₂ to glycine. With one more CO₂, glycine can be converted to serine and further to pyruvate and acetyl-CoA to yield more value-added products. (d) The serine cycle converts CO₂ and 5,10-methylenetetrahydrofolate (5,10-CH₂-THF), which can be derived from methane, methanol or formaldehyde, to acetyl-CoA. (e) The ribulose monophosphate cycle (RuMP) converts methane, methanol or formaldehyde to triose phosphate, which can feed into central carbon metabolism to produce cellular energy, biomass precursors, and bioproducts. Dark green boxes represent C1 substrates, the light green boxes represent potential products. Rate-limiting steps are underlined. ATP and redox cofactors are color-coded.

The WLP is an anoxic alternative to the CBB cycle for direct C2 synthesis (Figure 3b) [5]. In the WLP (also known as the reductive acetyl-CoA pathway), two CO₂ molecules are reduced in parallel via the methyl and carbonyl branches to form acetyl-CoA. The methyl branch of the WLP is one-carbon metabolism that is conserved across the kingdoms of life [5]. Methyl transfer has been suggested as the rate-limiting step of the WLP [19]. ATP is used for activating C1 in the methyl branch and a rate-limiting factor in autotrophy even though ATP is regenerated during acetate production. Controlled cofeeding of glucose to supply ATP stimulates CO₂ conversion into acetate [20]. In acetogens, Rnf and Nfn complexes contribute to energy conservation by chemiosmotic gradient buildup and electron bifurcation, respectively [21]. In Moorella thermoacetica, methylenetetrahydrofolate reductase (MTHFR), which converts methylene-THF to methyl-THF, is hypothesized to be electron bifurcating [22, 23]. Further studies into the energy-conserving mechanisms of the WLP would contribute to increasing the efficiency of input energy stored as energy-dense molecules [24]. The WLP serves as a steppingstone to converting CO₂ and CO into advanced bioproducts by producing acetate that heterotrophic microbes can utilize as sole carbon and energy sources. Two-stage processes converting syngas containing CO₂ and CO into acetate (e.g., using Clostridium aceticum and Moorella thermoacetica) and acetate into lipids (e.g., using Rhodotorula toruloides and Yarrowia lipolytica) have been demonstrated [20, 25, 26].

A partially reduced carbon formate is more water soluble compared to CO₂ and CO, albeit toxic at moderate concentrations (~150 mM) [27]. Formate can be reduced through the WLP methyl branch and the reductive glycine pathway (rGlyP). The rGlyP converts formate and CO₂ into glycine and with an additional formate into serine, which can be further converted into pyruvate (Figure 3c) [28]. In converting formate to glycine and serine, the rGlyP consumes ATP, NADPH, and NADH. While the reduction of 5,10-methenyl-THF to 5,10-methylene-THF is often rate-limiting in the rGlyP [28, 29], low ammonia concentrations has been shown to be a limiting factor in the rGlyP of Desulfovibrio desulfuricans [7]. The rGlyP is more ATP-efficient than the CBB cycle, it has proven efficient for C1 assimilation for cell growth and function in Pseudomonas putida and E. coli [29, 30].

Methane, methanol, and formaldehyde are challenging C1 substrates due to the gaseous nature of methane and the toxicity of formaldehyde. Life has still prevailed in evolving metabolic processes for detoxification and utilization of these substrates. Methane utilization in methanotrophs start by methyl transfer to C1 carriers or partial oxidation to methanol by using methane monooxygenase (MMO) [8, 31–33]. Mammals can dehydrogenate methanol to formaldehyde using alcohol dehydrogenase and further oxidize it to formate using aldehyde dehydrogenase [34]. Unicellular organisms like Pichia pastoris also use formaldehyde dehydrogenase to enable growth on methanol or formaldehyde. These relatively electron-rich carbons undergo both assimilative and dissimilative pathways for biomass production and energy generation [35]. To incorporate reduced C1 compounds into central carbon metabolism, type II methanotrophs use the serine cycle [36]. The serine cycle utilizes reduced C1 or formaldehyde and CO₂ to form a C2 acetyl group (Figure 3d) [37, 38]. A synthetic serine cycle has been introduced in E. coli for methanol assimilation [38]. However, its high energy cost of 2 ATP per cycle is costlier methylotrophy than that using the ribulose monophosphate (RuMP) cycle [39].

The RuMP cycle (Figure 3e) is an energy efficient methanol assimilation pathway with a Gibbs free energy change (ΔG’°) of −89.9 kJ/mol [40]. The RuMP cycle is popular for synthetic methylotrophy in E. coli because of minimal enzyme expression requirements [39]. E. coli and other organisms have been successfully engineered with the RuMP cycle [39, 41, 42]. Keller and colleagues identified 3-hexulose-6-phosphate synthase (HPS), which condenses formaldehyde and ribulose-5-phosphate (Ru5P) into hexulose-6-phosphate (Hu6P), as the limiting reaction [39]. Sanford and Woolston recognized that methanol assimilation through the RuMP cycle is thermodynamically constrained by formaldehyde buildup [43]. One drawback of this pathway is that when the three-carbon product, dihydroxyacetone phosphate (DHAP), is converted to acetyl-CoA for use in lipogenesis, one of the carbons is lost as CO₂. Thus, synthesizing products derived from C3 lower glycolytic intermediates is more desirable when using the RuMP cycle. The initial C1 compound and the desired final product should be paired using optimal pathways in terms of their carbon yield, ATP requirement, and heterologous gene expression.

Engineering Synthetic One-Carbon Metabolism

Integrating heterologous metabolic pathways into model organisms is a key objective of metabolic engineering. The benefit of using model organisms is that their well-characterized growth and metabolism makes them well suited for large-scale manufacturing [10]. By integrating natural one-carbon pathways into model organisms, researchers can get the best of both worlds. Chen and colleagues engineered a methylotrophic E. coli by introducing a modified RuMP cycle (Figure 4a) with three heterologous genes encoding methanol dehydrogenase (MDH), HPS, and 6-phospho-3-hexuloisomerase (PHI) [44]. To prevent formaldehyde toxicity, the engineered cells must keep the HPS reaction active and the RuMP cycle turning. The modified RuMP cycle minimizes the diversion of carbon flux away from HPS by using the Entner-Doudoroff (ED) pathway instead of PFK and fructose-1,6-bisphosphate aldolase (ALDO) for converting fructose-6-phosphate (F6P) into C3 lower glycolytic metabolites, one of which is the output of the RuMP cycle from three input C1 molecules. Using the ED pathway provides dual benefits: 1) the modified RuMP cycle produces pyruvate in fewer steps compared to the original RuMP cycle; and 2) the modified RuMP cycle does not require initial ATP investment [45]. By incorporating the modified RuMP cycle, Reiter and colleagues achieved synthetic methylotrophy in E. coli with a 4.3-hour doubling time and maximum culture density of 100 OD₆₀₀ in a fed-batch reactor that continuously fed and maintained methanol at >500 mM [46].

Figure 4. Synthetic C1 assimilation pathways integrated into host metabolism.

(a) A modified RuMP cycle introduced in E. coli leverages the ED pathway to supply C3 intermediates without ATP expenditure [44]. (b) Synthetic EuMP cycle converts formaldehyde to E4P with the introduction of EPS, LerI, and DerI enzymes. The cycle is completed by central carbon reactions. Adapted from [47]. (c) The synthetic homoserine cycle condenses formaldehyde with pyruvate to produce HOB and eventually acetyl-CoA. Adapted from [48]. (d) The HOB cycle is a synthetic alternative to one-carbon metabolism. Adapted from [49]. (e) Phosphate dependent formate to formaldehyde conversion can feed into formyl-CoA elongation reactions (FORCE) for synthesis of multicarbon units [50, 54]. Adapted from [54]. Dark green boxes represent C1 substrates, the light green boxes represent potential products. ATP and redox cofactors are color-coded. Segments of the pathways that are either modified, introduced in the organism or new-to-nature to create these cycles are highlighted in orange.

Designing novel pathways augments the toolset for C1 utilization. Wu and colleague designed the erythrulose monophosphate (EuMP) cycle for formaldehyde assimilation. Formaldehyde is condensed with DHAP to produce erythrulose-1-phosphate (Eu1P) and the product after three turns of the cycle is GAP (Figure 4b) [47]. The EuMP cycle is energetically efficient with just one ATP consumed per GAP produced. While it is just as energetically efficient as the RuMP cycle, the EuMP cycle includes more enzymatic steps.

The use of orthogonal molecules mitigates the potential issue of substrate competition between native and engineered pathways. The synthetic homoserine cycle for methanol assimilation condenses formaldehyde with pyruvate to produce 4-hydroxy-2-oxobutanoic acid (HOB), which can be converted to homoserine (Figure 4c) [48]. Similarly, a synthetic 4-hydroxy-2-oxobutanoic acid (HOB)-dependent one-carbon metabolism has been introduced in E. coli as an alternative to the serine cycle (Figure 4d) [49]. This pathway relies on two novel reactions: the transamination of L-homoserine and transfer of one carbon unit from HOB to THF with release of pyruvate. The strain assimilates CO₂ without relying on endogenous metabolism. Chou and colleagues designed an orthogonal metabolic framework in E. coli for creating multi-carbon products directly from C1 substrates using formyl-CoA elongation reactions (FORCE) so that the assimilation does not compete with the host metabolism [50]. Intermediates from these pathways can be readily transformed to several value-added products (2-hydroxy acids, aldoses, diols, polyols, carboxylic acids and alcohols) as well as biomass precursors. Thermodynamic and stoichiometric analyses identified ATP consumption, the NADH/NAD⁺ ratio, and formate concentration as major constraints to the max-min driving force (MDF) of the FORCE pathway [51]. The FORCE pathway was introduced in E. coli and tested in a co-culture system, in which one strain could convert formate, methanol, and formaldehyde to glycolate while the other could consume glycolate for growth.

Engineering new enzymes benefits C1 utilization on an individual reaction level and helps conceive new pathways. The condensation of formyl-CoA and formaldehyde is a major bottleneck in the FORCE pathway [50, 52]. The oxalyl-CoA decarboxylase from Methylobacterium extorquens was mutated to a glycolyl-CoA synthase (MeOXC4), which allows for C₁-C₁ condensation of formyl-CoA and formaldehyde. MeOXC4 had 40-fold more catalytic efficiency compared to all other known natural and synthetic enzymes for C₁-C₁ condensation at the time [53]. Since then, the variants of 2-hydroxy-CoA synthase (HACS) with improved activities have been identified [52]. Nattermann and colleagues also introduced a two-step cascade for phosphate-dependent conversion of formate to formaldehyde [54]. The new-to-nature enzyme formyl phosphate reductase (FPR) boasts a 300-fold shift in formyl phosphate specificity. Combining FPR with the FORCE pathway [50] allows C2 production with formate as the sole carbon source (Figure 4e).

Powering Metabolism to Attain the Circular Economy

A common requirement for operating one-carbon metabolism is energy in the form of reducing power and high-energy phosphates. For example, acetogenesis requires 2 CO₂ and 8 electrons with the redox potential of CO₂/acetate of −0.28 V versus standard hydrogen electrode (SHE) [55]. In biological systems, ATP is required for driving thermodynamically unfavorable reactions and running cellular housekeeping processes. Substrate cofeeding combined with systems and synthetic biology tools increases the efficiency of C1 conversion by better affording cellular energy. Furthermore, electrochemical cells and light-harvesting materials provide alternative means to power bioenergetic pathways. Utilization of diversified energy sources will engender a resilient circular bioeconomy.

Balanced Cofactor Generation Powers Metabolism

Efficient bioproduct synthesis using C1 compounds necessitates a balanced supply of carbon backbones, ATP, and reducing equivalents (Figure 5a). Canonical redox pairs are ubiquitously employed across metabolism (Box 1). Healthy metabolic function hinges on their availability [56]. Synthetic biology efforts have sought to improve and balance cofactor generation [57, 58], favor one cofactor over another [59], and introduce non-canonical redox cofactors to minimize loss of electrons to competing pathways [56]. One approach to overcoming imbalanced cofactor production is substrate cofeeding, which activates different metabolic pathways without genetic engineering [60]. Feeding C1 substrates with sugars such as glucose and fructose has been shown to improve energy efficiency and carbon yield. Doping gas-fermenting M. thermoacetica with glucose streamlines ATP generation to accelerate cellular metabolism and CO₂ reduction [20]. Limiting glucose supply limits its accumulation in the culture and overcomes catabolite repression in the mixed substrate cofeeding while activating glycolytic ATP generation. A mixture of CO₂, CO, and H₂ can be fed to acetogens to power the WLP. In the WLP, CO₂ and CO provide carbon backbone whereas the oxidation of CO and H₂ provide reducing equivalents to create NAD(P)H and reduced ferredoxin. Mixotrophic growth on C1 compounds, sugars, and other nutrients promotes fast metabolism and bioproduct synthesis [61–63]. Another approach to overcome redox cofactor limitations is to utilize alternative cofactors, which has been implemented in E. coli to support cell growth and channel reducing power from glucose to levodione through nicotinamide mononucleotide (NMN) rather than NADPH [64]. The cofactor specificity of enzymes for NADH and NADPH may be swapped. The mevalonate pathway for isoprenoid synthesis uses either NADPH or NADH depending on the organism. In Halomonas bluephagenis, NADH-dependent strains produced more mevalonate because NADH is more abundant in the cell [65]. In engineered E. coli methylotrophs, the MDH reaction which converts methanol to formaldehyde is inhibited by high NADH concentrations. To overcome this, researchers have sought to couple methanol oxidation to an artificial NADH consumption pathway [66]. These synthetic biology approaches to boost the availability and efficient use of redox cofactors help address challenges in C1 substrate utilization.

Figure 5. Chemical, electric, and solar energy powers metabolism.

(a) Balanced supply of carbons, reducing power, and ATP via metabolic engineering and substrate cofeeding accelerates C1 utilization. (b) Electrochemistry-powered bioenergetic pathway. Carrier molecules (e.g., H₂ and syngas) transfer reducing power from cathode to bacteria and support C1 conversion and autotrophy. (c) Light-powered bioenergetic pathway. Light-harvesting materials (e.g., quantum dots) harness solar power and transfer reducing equivalent to microbes through cell-material interface, creating an artificial photosynthetic system.

Box 1. Efficient energy utilization using disparate redox cofactors.

Redox cofactors mediate electron transfers and couple anabolic and catabolic processes in metabolism. Each redox pair has a unique redox potential. Enzymatic steps in C1 assimilation pathways utilize different redox cofactors (Figure 3). Nicotinamide cofactors (NAD and NADP and their reduced equivalents NADH and NADPH) both have reduction potential E₀^’=–320mV however their redox potentials differ due to the availability of their reduced and oxidized pools [56]. The NAD/NADH ratio is kept very high by shuttling electrons to the electron transport chain and is used in oxidation reactions. The NADP/NADPH ratio is kept low due to the role of NADPH as a reducing power in biosynthesis. NADPH is produced by the oxidative pentose phosphate pathway and malic enzyme as well as one-carbon metabolism and isocitrate dehydrogenase in some organisms. Ferredoxins, on the other hand, are iron-sulfur proteins with reduction potential E₀^’~–420mV [87, 88]. In contrast to nicotinamide cofactors which mediated two electron transfers at time, ferredoxins typically mediate a single electron transfer. Cells have ferredoxins with different Fe-S clusters, each tuned for specific reactions. We have shown the most reported cofactors for each reaction (Figure 3), however different species may possess different enzymes to carry out the same reaction using different cofactors. A more detailed view of these pathways can be found on MetaCyc (MetaCyc.org), where for example the methylene tetrahydrofolate reaction (MTHFR) in the WLP shows three possible cofactor pairs for the reduction reaction and different possible enzymes. Interestingly, MTHFR reaction in M. thermoacetica is thought to operate electron bifurcation to minimize overpotential. Electron bifurcation transfers electrons from one source to two disparate electron carriers, one with more negative reduction potential for overcoming “difficult” energy barriers (e.g., in CO₂ reduction) and the other with less negative reduction potential to limit wasting free energy [88]. Incorporating electron bifurcation in anaerobic digestion has been shown to increase ATP generation and methane production under substrate limitation [89]. Introducing a flavin-based electron bifurcation in E. coli led up to a 4-fold increase in H₂ and succinate production, 7-fold increase in ATP accumulation and superior cell-fitness [90].

Electrochemistry Powers Metabolism

In electrochemistry-assisted microbial CO₂ fixation, cathodes power CO₂ fixation by transferring reducing equivalents such as H₂ to cells (Figure 5b). A cobalt-phosphorus alloy cathode has been used to produce H₂ to enhance CO₂ fixation process of Ralstonia eutropha [67]. Deutzmann and colleagues designed a hybrid biochemical CO₂ fixation system including a biocathode consisting of a graphite electrode and Desulfopila corrodens strain IS4 [68]. H₂ production speed on the biocathode exceeds that of platinum electrode, a widely used inorganic catalyst. The feasibility of the biocathode-driven CO₂ fixation has been demonstrated using a methanogen Methanococcus maripaludis and an acetogen Acetobacterium woodii for methane and acetate production. A photovoltaic device has been used to electrochemically reduce CO₂ and H₂O into syngas which was further transformed into butanol and hexanol using Clostridium autoethanogenum [69]. Remarkably, this hybrid process featured faradaic efficiency as high as 80–100% and a lower energy cost compared to a chemical synthesis route.

Electrochemistry-assisted bacterial CO₂ fixation has lower energy costs and higher efficiency than photosynthesis, but maximum CO₂ conversion efficiency is limited by the low solubility of electron mediators (e.g., H₂). To overcome the obstacle, perfluorocarbon (PFC) nano-emulsion has been used as a H₂ carrier to promote acetate production by Sporomusa ovata [70]. Due to PFC’s excellent H₂ solubility, addition of nano-emulsion increases the throughput of CO₂ reduction into acetate by as much as 190%. Claassens and colleagues found that formate and methanol can work as electron mediators to supply reducing power from electrodes to microbes at a high energetic efficiency [71]. Despite the simple nature of acetate, electrochemistry-driven CO₂ reduction into C2 products is beneficial since heterotrophic microbes can utilize the C2 compounds as the sole carbon and energy source [72].

Besides providing reducing power, electrochemistry enables fine tuning of microenvironments. Liu and colleagues utilized Pt-coated Si wire array to achieve CO₂-to-acetate transformation of a strict anaerobe S. ovata under aerobic conditions [73]. With solar energy input, Pt catalyzes O₂ reduction by photocurrent in the local environment, leading to microscopic anoxic conditions to support the S. ovata culture. Similarly, electrochemical O₂ microscopic gradients have been used to accommodate N₂-fixing bacteria [74] for accelerated nitrogen assimilation. A unique benefit of introducing electrochemistry to bioprocesses is the control it offers. By adjusting applied voltage and electrode morphology, energy can be efficiently delivered to cells. Machine learning has proven useful in tuning these variables to create suitable microenvironments for microbial C1 conversion to afford promising avenues for biomass and bioproduct synthesis [75].

Light-Harvesting Material Powers Metabolism

Light-harvesting materials can transduce solar energy into reducing power for microbes (Figure 5c). Material-bacteria interfaces thus give rise to artificial photosynthetic systems. One commonly used light-harvesting material is Si wire array. A Si-nanowire-bacteria hybrid was the first example of direct interface between CO₂-reducing S. ovata and semiconductors allowing microbial photoelectrosynthesis for CO₂-to-acetate transformation [73]. Adding on an N₂-fixing bacterium Rhodopseudomonas palustris to the nanowire-bacteria system, the co-culture system achieved a solar-to-chemical efficiency of 0.51% for nitrogenous biomass [76]. A hybrid of quantum dots and bacteria also creates an artificial photosynthesis system [77]. CdS quantum dots can capture sunlight and produce reducing equivalents to operate the WLP in M. thermoacetica. CdTe quantum dots have been interfaced as light harvesters on Xanthobacter autotrophicus for simultaneous CO₂ and N₂ fixation [78]. High photophysical charge-transfer kinetics between quantum dots and microbes enables CO₂ fixation at an internal quantum efficiency of ~47%, close to the biochemical limits at 46.1%. Semiconductors La- and Rh-doped SrTiO₃ and Mo-doped BiVO₄ have been used as a photocatalyst sheet for absorbing solar energy for water splitting, and providing S. ovata with H₂ for the CO₂-to-acetate conversion [79]. An organic semiconductor (e.g., a heterojunction of perylene diimide derivative and poly(fluorene-co-phenylene)) has been used to generate photoexcited electrons for nonphotosynthetic bacteria M. thermoacetica to reduce CO₂ into acetate [80]. Thus, light-harvesting materials expand the repertoire of renewable energy sources for microbial C1 conversion.

Toward hybrid powering of metabolism

The ability of microbes to harness multiple energy sources promotes bioproduct synthesis, rain or shine. The versatile energy utilization can be accomplished on an individual culture level, in which an isogenic or heterogeneous population of cells have access to chemical, electrical, and light energy, or in distributed cultures that work in concert to upcycle C1 compounds. The hybrid powering strategy may combine the benefits of nutrient cofeeding that activates multiple parts of metabolism for advanced bioproduct synthesis, electrochemistry that confers excellent controllability, and abiological material that harvests and transduces plentiful solar energy. We envision interdisciplinary projects coordinating to construct a “power grid” for bioeconomy.

Concluding Remarks: Bringing Elements to Life

Engineering and powering metabolism through cofactor balancing and green energy provide a blueprint for repurposing of C1 compounds to value-added products. Central carbon metabolism acts as a rendezvous for one-carbon metabolism and biosynthetic metabolism. Matchmaking between C1 substrates and advanced bioproducts requires contiguous enzymatic reaction steps that bridge them and energy metabolism that powers the assimilative and biosynthetic pathways. Thus, systems-level biochemical and energetic considerations help conceive scientifically and commercially viable C1-bioconversion processes.

Integration of analytical, biochemical, and computational tools provides a holistic understanding of one-carbon metabolism from thermodynamic and kinetic perspectives (see Outstanding Questions). In heterotrophic organisms, mass spectrometry, ¹³C stable isotope tracing, and mathematical optimization have imparted flux information (i.e., rates at which cells operate metabolic pathways) based on steady-state isotope labeling patterns in metabolites that imprint metabolic activities. In autotrophs, ¹³C tracing from C1 compounds is more difficult as the steady-state ¹³C-labeling state comprises uniformly labeled metabolites, and thus dynamic labeling strategies are preferred [81–83]. A quantitative understanding of one-carbon metabolism flux and regulation will engender more rational metabolic engineering and synthetic biology endeavors for C1 upcycling.

Outstanding Questions.

• Cells are made of several major elements including carbon and nitrogen. Thus, carbon and nitrogen metabolism must be coordinated for efficient biomass and bioproduct synthesis. How can we better understand the regulation and coordination of carbon and nitrogen metabolism? How can the resulting knowledge help repurpose nitrogen waste simultaneously with carbon waste?

• Problems at hand require interdisciplinary approaches. How can biochemical, genetic, and computational tools be integrated to facilitate efficient energy transduction, biosynthesis, and their coupling?

• What are technoeconomically feasible C1-to-product conversion processes?

• How can we translate laboratory knowledge to industrial scale bioprocesses to engender a positive global impact?

From a technoeconomic perspective, additional considerations of scale-up, market sizing, and process design are needed. Only when the biological process yields cost-effective products at scale compared to existing chemical processes would value be created. For example, LanzaTech has successfully commercialized a gas-fermentation process using syngas from steel mills to produce ethanol and acetone using acetogenic Clostridia [84]. Others have worked towards commercial syngas fermentation; however, challenges related to scale-up, gas-to-liquid mass transfer, and costs linger [85]. While powering one-carbon metabolism to convert C1 substrates into C2 and C3 intermediate products acts as a store of value, attaining C4+ products can drive value creation. Commercialization of C1-derived jet fuels and advanced bioproducts such as polyketides and terpenoids would yield environmentally and economically productive outcomes.

In addition to carbon, nitrogen constitutes a large portion of the biogeochemical cycle. Nitrogen-fixing bacteria as well as the Haber-Bosch process produce ammonia from N₂. Nitrate and nitrite from runoff wastewater may also be converted to ammonia using electrochemical processes [86]. Integrative research into carbon and nitrogen metabolism will bring us closer to a circular economy (see Outstanding Questions).