Recent advancements in microbial carbon dioxide fixation: metabolic engineering strategies

Abstract

Global warming driven by rising CO₂ emissions has spurred intense interest in biological carbon fixation as a sustainable alternative to physicochemical approaches. This review explores natural, engineering, and synthetic CO₂ fixation strategies in both microbial and cell-free systems. We first outline canonical natural pathways, then highlight recent metabolic engineering approaches that enhance flux through native enzymes, optimize cofactor balances, and leverage CRISPR- and evolution-based tools to improve growth on CO₂ or mixed C1 substrates. Next, we describe emerging synthetic routes, as well as electrochemical–microbial hybrid systems that couple renewable electricity with biocatalysis. We then discuss downstream engineering tactics for metabolic-flux redirection, energy-supply augmentation, regulatory-gene editing, and enzyme-design via directed evolution to overcome kinetic and thermodynamic barriers. High-density cultivation, omics-guided optimization, and non-traditional hosts (acetogens, sulfate reducers, methanotrophs, methylotrophs, and yeasts) are surveyed for their industrial potential. Finally, we evaluate integration with waste-gas streams, direct-air-capture platforms and biomass conversion, consider techno-economic and scale-up challenges, and identify opportunities for artificial-intelligence driven pathway design. By uniting advances across biology, chemistry, and engineering, this review charts a roadmap toward efficient and scalable CO₂-to-chemical bioproduction. Future progress in microbial CO₂ fixation is expected to rely on the development of more efficient CO₂-fixing enzymes and metabolic pathways, together with the incorporation of electro-biotechnology and optimized bioprocesses to enhance scalability and carbon conversion efficiency.

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Green foundation

1. As metabolic engineering of microbes emerges as a prominent combative force to mitigate greenhouse gas (GHG) emissions such as CO₂, we provide a rundown of natural and synthetic pathways utilized by microbes to accomplish this task. Various strategies for engineering and optimizing CO₂-fixing microbes have been discussed.

2. Utilizing microbes as a green platform not only to reduce atmospheric CO₂ but also to produce valuable compounds holds promise, as it can turn 37.8 Gt of pollutants into a potential industrial feedstock. We highlight techniques that bring the synthetic biology, process engineering, electrochemistry, and computational domains together to improve this technology in the future.

3. Although much work has been focused on industrializing the concept, the scalability of this technology remains a challenge as of now. The review summarizes various industrial bottlenecks and addresses them with multidisciplinary strategies.

1. Introduction

Global warming is one of the most pressing challenges facing the world today, with carbon dioxide (CO₂) being a major contributor. Each year, CO₂ emissions continue to rise at an alarming rate, reaching 41.6 billion tons in 2024.¹ To address this challenge, various strategies have been implemented, including reforestation, afforestation, and soil carbon sequestration, to help reduce CO₂ levels and mitigate global warming.² Microorganism-based approaches have also been explored, enabling the fixation of CO₂ into products such as biofuels, bioplastics, and other industrially important compounds.^3,4

Microorganisms naturally use CO₂ fixation pathways, which can be enhanced through metabolic engineering strategies to improve efficiency. Enzyme engineering has significantly contributed to enhancing CO₂ fixation efficiency, as demonstrated by both direct and indirect modifications of the ribulose-1,5-bisphosphate carboxylase/oxygenase (RubisCO) enzyme.^5,6 Additionally, synthetic pathways have been designed to channel CO₂ into the production of valuable compounds.⁷ Recent advancements, particularly the integration of artificial intelligence, have further accelerated the optimization of metabolic pathways and enzyme engineering, enabling more efficient and precise CO₂ fixation strategies.

Despite recent advancements, major challenges persist, most notably the low efficiency of carbon utilization and limited bioproduction. The current capacity of CO₂ fixation technologies remains insufficient to meet global sustainability demands. To address these gaps, this review examines natural, engineered, and synthetic metabolic pathways for CO₂ fixation. It explores the key enzymes and cofactors involved, metabolic pathway kinetics, and electron transport mechanisms. Additionally, we discuss recent advancements in sustainable bioproduction from CO₂, along with the expanding role of artificial intelligence in accelerating progress in this field.

2. Microbial CO₂ fixation pathways and chemical conversion

2.1. Natural CO₂ fixation pathways

Natural CO₂ fixation pathways are fundamental to converting atmospheric CO₂ into organic biomass, sustaining life, and maintaining global carbon cycles. The CBB cycle, predominant in plants, algae, and cyanobacteria, relies on RubisCO to catalyze CO₂ assimilation.⁸ In contrast, anaerobic bacteria and archaea utilize alternative pathways tailored to their environmental constraints. For instance, the reductive tricarboxylic acid (rTCA) cycle fixes CO₂ by running the oxidative TCA cycle in reverse, while the 3-hydroxypropionate bicycle, found in Chloroflexus bacteria, and the Wood–Ljungdahl pathway (WLP), used by acetogens and methanogens, function efficiently under anaerobic conditions (Fig. 1).

Fig. 1 The illustration of multiple natural CO₂ fixation pathways into central carbon metabolism. The pathways include the Calvin–Benson–Bassham (CBB) cycle, the Wood–Ljungdahl pathway, the 3-hydroxypropionate (3HP) bicycle, the 3-hydroxypropionate/4-hydroxybutyrate (DC/4HB) cycle, and the reductive tricarboxylic acid (rTCA) cycle. Carbon fixation steps are marked by the incorporation of CO₂ or HCO₃⁻, catalyzed by key enzymes such as ribulose-1,5-bisphosphate carboxylase/oxygenase (RubisCO), acetyl-CoA carboxylase (ACC), carbon monoxide dehydrogenase/acetyl-CoA synthase (CODH/ACS), phosphoenolpyruvate synthase and carboxylase (Ps & Ppc), isocitrate dehydrogenase (ICDH), and propionyl-CoA carboxylase (Pcc). The figure also highlights metabolic energy requirements via ATP consumption and redox balancing through NADPH use. Abbreviations of metabolites are as follows: R5P – ribulose 5-phosphate; F6P – fructose 6-phosphate; G3P – glyceraldehyde 3-phosphate; PYR – pyruvate; Ac-CoA – acetyl-CoA; M-CoFeS – methyl-CoFeS; MTFH – methenyl-tetrahydrofolate; MeTHF – methyl-tetrahydrofolate; 10-FTHF – 10-formyl-tetrahydrofolate; FOR – formate; Mal-CoA – malonyl-CoA; MeC4-CoA – methylsuccinyl-CoA; 3HP – 3-hydroxypropionate; 3Pp-CoA, 3-propionyl-CoA; mm-CoA – methylmalonyl-CoA; OAA – oxaloacetate; MAL – malate; FUM – fumarate; SUC – succinate; Suc-CoA – succinyl-CoA; CIT – citrate; 2OG – 2-oxoglutarate (α-ketoglutarate); Cr-CoA – crotonyl-CoA; PEP – phosphoenolpyruvate; 4-HB-CoA – 4-hydroxybutyryl-CoA; 4-HB – 4-hydroxybutyrate; and Ssa – succinic semialdehyde.

Expanding this metabolic diversity, certain archaea in extreme environments, such as thermoacidophilic archaea (Sulfolobus, Acidianus, and Metallosphaera), employ the 3-hydroxypropionate/4-hydroxybutyrate (3-HP/4-HB) cycle, which merges 3-HP and 4-HB intermediates to optimize CO₂ fixation. Similarly, the dicarboxylate/4-hydroxybutyrate (DC/4-HB) cycle, used by anaerobic archaea like Ignicoccus, integrates dicarboxylate and 4-HB metabolism to fix CO₂ with minimal ATP consumption, making it energy-efficient for nutrient-limited settings.⁹

2.2. Synthetic CO₂ fixation pathways

Synthetic CO₂ fixation pathways represent innovative biochemical routes designed to enhance carbon capture and utilization beyond natural pathways like the Calvin cycle. The CETCH cycle (crotonyl-CoA/ethylmalonyl-CoA/hydroxybutyryl-CoA) is a notable example, combining enzymes from diverse organisms to achieve higher ATP efficiency than those in natural cycles.¹⁰ An engineered variant of the serine cycle, incorporating an alternative CO₂ fixation pathway via pyruvate, has been successfully demonstrated in E. coli.¹¹ The ASAP (artificial starch anabolic pathway), a chemical–biochemical hybrid pathway for starch synthesis from CO₂ and hydrogen, has been established in a cell-free system.¹² Similarly, formolase catalyzes the formation of a carbon–carbon bond by condensing three molecules of formaldehyde into one molecule of the three-carbon compound dihydroxyacetone.¹³ The SACA (synthetic acetyl-CoA) pathway, a synthetic, ATP-independent pathway, has been developed to produce acetyl-CoA from one-carbon substrates such as formaldehyde.¹⁴ These pathways highlight the potential of synthetic biology to reimagine carbon fixation, offering scalable solutions for sustainable bioproduction and climate change mitigation. Fig. 2 illustrates engineered and synthetic variants of CO₂ fixation, and Fig. 3 compares their energy demands and engineering complexity. Table 1 complements these figures by summarizing core CO₂ fixation pathway characteristics.

Fig. 2 A network of engineered CO₂ assimilation pathways integrated into central metabolism. Highlighted pathways include the GED (Gnd-Edd-Eda) pathway, the reductive glycine (rGly) pathway, the modified serine cycle, the reductive glyoxylate and pyruvate synthesis (rGPS) cycle, the TaCo (tartronyl-CoA) pathway, the CETCH cycle, the THETA cycle, and the HOPAC cycle. Key enzymatic steps for carbon fixation and C1 assimilation are labeled, including 6-phosphogluconate dehydrogenase (Gnd), Entner–Doudoroff enzymes (Edd, Eda), serine hydroxymethyltransferase (SHMT), the glycine cleavage system and tetrahydrofolate pathway (Gcv/THP), methylene-THF dehydrogenase (MtdA), formyl-THF cyclohydrolase (Fch) and formyl-THF ligase (Fti), phosphoenolpyruvate carboxylase (Ppc), the glycine cleavage system (GCS), glyoxylate carboxylase (GCC), crotonyl-CoA carboxylase/reductase (Ccr), malonyl-CoA synthase (MAS), and propionyl-CoA carboxylase (Pcc). Abbreviations of metabolites are as follows: R5P – ribulose 5-phosphate; F6P – fructose 6-phosphate; G3P – glyceraldehyde 3-phosphate; 6PG – 6-phosphogluconate; KGPG – 2-keto-3-deoxy-6-phosphogluconate; PYR – pyruvate; SER – serine; PEP – phosphoenolpyruvate; OAA – oxaloacetate; MAL – malate; MTFH – methenyl-tetrahydrofolate; MeTHF – methyl-tetrahydrofolate; 10-FTHF – 10-formyl-tetrahydrofolate; FOR – formate; Mal-CoA – malonyl-CoA; Ma-CoA – malyl-CoA; Cr-CoA – crotonyl-CoA; Ace-CoA – acetyl-CoA; Gly – glycine; Go – glyoxylate; Ga – glycolate; Gc – glycerate; Gl-CoA – glyceryl-CoA; Tt-CoA – tartronyl-CoA; 4HB – 4-hydroxybutyrate; Em-CoA – ethylmalonyl-CoA; Prp-CoA – propionyl-CoA; Acr-CoA – acryloyl-CoA; Me-CoA – methylmalonyl-CoA; Suc – succinate; Suc-CoA – succinyl-CoA; Fum-CoA – fumaroyl-CoA; Ala-CoA – alanoyl-CoA; Ala – alanine; 3HP – 3-hydroxypropionate; and 3HP-CoA – 3-hydroxypropionyl-CoA.

Fig. 3 Comparison of natural and synthetic CO₂ fixation pathways. Natural pathways (e.g., CBB cycle, 3HP bicycle, rTCA cycle, WLP, 4HB pathway) are typically hosted in autotrophs such as C. necator and cyanobacteria, offering robustness but posing challenges for genetic engineering. Synthetic pathways (e.g., CORE, CETCH, rGly, ASAP, HOPAC, SACA, formolase) have been implemented in tractable hosts like E. coli and S. cerevisiae, enabling easier engineering and product diversification. Trade-offs between energy/redox requirements, host choice, and engineering complexity determine their suitability for applications ranging from biofuels to bioplastics.

Table 1 Comparison of natural and synthetic carbon dioxide fixation pathways in terms of various parameters

Pathway	No. of enzymes involved	Key enzymes involved in the pathway	Oxygen sensitivity	End products	Energy efficiency (per carbon)	Ref.
Natural pathways
Calvin–Benson–Bassham (CBB) cycle	11	RubisCO	Aerobic	Glyceraldehyde-3-phosphate	3 ATP and 2 NAD(P)H	25
Reductive tricarboxylic acid (rTCA)	8	2-Oxoglutarate synthase, isocitrate, dehydrogenase, fumarate reductase	Anaerobic	Acetyl-CoA	1 ATP and 2 NAD(P)H	26
Wood–Ljungdahl (WL) pathway	8–10	NAD-independent formate dehydrogenase, acetyl-CoA synthase-CO dehydrogenase	Anaerobic	Acetyl-CoA	0.5 ATP and 2 NAD(P)H	27
3-Hydroxypropionate bicycle	13	Acetyl-CoA carboxylase, propionyl-CoA carboxylase	Aerobic	Pyruvate	1.67 ATP and 2 NAD(P)H	28
3-Hydroxypropionate/4-hydroxybutyrate (3-HP/4-HB) cycle	15	Acetyl-CoA carboxylase, propionyl-CoA carboxylase	Aerobic	Acetyl-CoA	2 ATP and 2 NAD(P)H	29
Dicarboxylate/4-hydroxybutyrate (DC/4-HB) cycle	14	Pyruvate synthase, PEP carboxylase	Anaerobic	Acetyl-CoA	1.5 ATP and 2 NAD(P)H	30
Synthetic pathways
Crotonyl-CoA/ethyl malonyl-CoA/hydroxybutyryl-CoA (CETCH) pathway (version 5.4)	17	CoA-dependent carboxylase	Aerobic	Glyoxylate	0.5 ATP and 2 NAD(P)H	10
Reductive glycine pathway	8,9	Formate dehydrogenase, glycine cleavage system	Aerobic	Pyruvate	2 ATP and 3NAD(P)H	25
Artificial starch anabolic pathway (ASAP) (version 3.0 and 3.1)	10 (plus CO2 to methanol chemical reaction)	Formolase, Dihydroxyacetone kinase, ADP-glucose pyrophosphorylase	N/A	Amylose/amylopectin	2 ATP per C6 (~0.33 ATP /CO2)	12
Synthetic acetyl Co-A (SACA) pathway	3 (from formaldehyde; +2 if from CO2)	Glycolaldehyde synthase	Aerobic and anaerobic	Acetyl-CoA	No ATP or NAD(P)H required from formaldehyde. Starting from CO2 requires 2 NAD(P)H	14
HydrOxyPropionyl-CoA/Acrylyl-CoA (HOPAC) cycle (version 4.0)	11	Acetyl-CoA carboxylase, propionyl-CoA carboxylase, cinnamoyl-CoA reductase	Aerobic	Glyoxylate	1ATP, 1.5 NAD(P)H	31
Formolase pathway	5	Acetyl-CoA synthase, acetaldehyde dehydrogenase, formolase (FLS)	Aerobic and anaerobic	Dihydroxyacetone	2 ATP and 2 NAD(P)H	13
CO2 reduction (CORE) cycle (CoA-transferase variant)	6	β-Keto acid cleavage enzyme (BKACE)	Aerobic	Acetyl-CoA	1 ATP and 1 NAD(P)H	32

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2.3. Chemical conversion of CO₂

Electrochemical CO₂ conversion utilizes renewable electricity to reduce CO₂ into value-added chemicals (e.g., hydrocarbons, alcohols, syngas) which can then be harnessed by microorganisms as feedstocks for the sustainable production of biofuels and bioplastics. Central to this process are catalysts (e.g., copper, zinc, or molecular complexes) that lower the energy barrier for CO₂ activation and steer selectivity toward desired products. Key pathways include the reduction of CO₂ to CO pathway (via CO₂-to-CO electrocatalysts like Cu, Sb), the formic acid and methane pathway (using Cu/Zn), and the multi-carbon product pathway (e.g., ethylene/ethanol on Cu-based catalysts).^15–17 A representative biohybrid strategy demonstrated gram-scale PHB production by integrating electrochemical CO₂ reduction with microbial fermentation. In this system, CO₂ was electrochemically reduced to formate using Sn-based gas diffusion electrodes (GDEs), which then served as the carbon source for C. necator. The cells accumulated PHB up to 83% of dry cell weight, and with a 4 cm² Sn-GDE the process yielded 1.38 g of PHB, demonstrating the feasibility of direct CO₂-to-bioplastic conversion.¹⁸ Beyond polymers, engineered C. necator LH74D has been applied for electrobiosynthesis of liquid fuels, including isobutanol and 3-methyl-1-butanol. In this integrated electro-microbial system, CO₂ was first reduced to formate, which functioned as both a carbon and an energy source. Operating within a single reactor, the platform produced over 140 mg L⁻¹ of biofuels directly from electricity and CO₂, illustrating the potential of electro-microbial coupling for sustainable fuel synthesis.¹⁹

Beyond electrochemical reduction, chemical CO₂ conversion also encompasses thermochemical and photochemical approaches. Thermochemical routes, such as catalytic hydrogenation using metals like Ru, Ni, or Fe, convert CO₂ into methanol, dimethyl ether, or hydrocarbons under high temperature and pressure.^20,21 Photocatalytic systems, often employing TiO₂ or metal–organic frameworks (MOFs), enable solar-driven CO₂ reduction to fuels such as methane and methanol.²² In one study, photocatalyst-assisted PHB production was demonstrated using C. necator in combination with cadmium sulfide (CdS) nanorods. Under light irradiation, the CdS nanorods transferred photoexcited charges to C. necator, facilitating CO₂ reduction and channeling the fixed carbon into PHB biosynthesis. This hybrid system achieved a yield of 28 mg of PHB from CO₂ within 48 hours.²³ In another study, a biohybrid system was developed using the acetogen Sporomusa ovata coupled with a photocatalyst sheet to overcome the challenge of external electricity input. The photocatalyst sheet was composed of two particulate semiconductors La and Rh co-doped SrTiO₃ (SrTiO₃:La,Rh) and Mo-doped BiVO₄ (BiVO₄:Mo). Under sunlight, CO₂, and H₂O, the system simultaneously oxidized water to generate O₂ while transferring electrons and H₂ to S. ovata, which selectively converted CO₂ into acetate (CH₃COO⁻). This hybrid approach achieved a solar-to-acetate conversion efficiency of 0.7% under ambient conditions.²⁴ Taken together, these integrated chemical–biological systems represent a promising route toward carbon-neutral manufacturing of sustainable materials and energy carriers.

3. Recent advances in genetic and system engineering for CO₂ fixation

Microbial carbon dioxide fixation can be improved through a combination of genetic, metabolic, and process-level strategies (Fig. 4). Techniques such as enzyme engineering and CRISPR-based gene editing allow for precise modifications to pathways, enhancing fixation efficiency. Optimizing metabolic flux, electron donors, and energy balance ensures effective CO₂ assimilation by maintaining resource equilibrium. Adaptive laboratory evolution and omics-based approaches help identify and develop high-performance strains. Additionally, optimizing high cell-density cultures enhances carbon capture efficiency in industrial settings. These integrated strategies collectively contribute to advancing microbial CO₂ fixation.

Fig. 4 The framework of key optimization strategies for enhancing microbial carbon dioxide fixation, including enzyme engineering, CRISPR-based editing, metabolic flux and electron donor optimization, adaptive laboratory evolution, omics-driven approaches, high cell-density culture optimization, and energy optimization.

3.1 Metabolic flux optimization

Microbial CO₂ fixation is vital for reducing carbon emissions but is often limited by inefficient enzymes or metabolic pathways. Enhancing the expression of native carbon fixation genes or introducing more effective heterologous genes can address this limitation. For instance, a heterotrophic E. coli strain engineered to express korA and korB from Chlorobaculum tepidum encoding ferredoxin oxidoreductase demonstrated enhanced incorporation of CO₂ in the rTCA cycle metabolites such as citrate, pyruvate, α-ketoglutarate, and succinate.³³ Enhancing metabolic flux by supporting the function of key enzymes can help overcome carbon fixation inefficiencies. In autotrophs, the primary limitation is the low carbon capture efficiency of RubisCO. Introducing exogenous carbonic anhydrase, which converts CO₂ to bicarbonate, increases carbon availability to RubisCO. This has improved the RubisCO efficiency in microalgae like Chlorella sp. by boosting enzyme levels by up to 25% and reducing oxygenation rates.³⁴ It has been shown that increases in concentrations of succinate and malate also positively influence the rate efficiency of carbon dioxide assimilation by directing the carbon flux to efficient pathways and reducing carbon loss.³⁴ Heterologous expression of phosphoenolpyruvate carboxylase and phosphoenolpyruvate carboxykinase in a type I methanotroph increased CO₂-dependent succinate production by 46% compared to the wild type.³⁵

3.1.1. Optimization of energy supply. The high energy requirements of the associated enzymes, which vary depending on the process, present a significant obstacle to biological CO₂ fixation. For instance, the CBB cycle depends on ATP and NADPH, while the WL pathway in Clostridium and Acetobacterium uses reducing agents like H₂ or formate.³⁶ In E. coli, the expression of korA and korB enabled electron transfer analogous to the rTCA cycle, which in organisms such as Desulfobacter and Chlorobaculum depends on ferredoxin-linked enzymes with high reducing power requirements.³⁷ Thus, boosting the availability of cofactors such as ATP, NAD(P)H, or noncanonical redox cofactors (NCRCs) is a promising strategy to overcome these energy constraints.³⁸ Since these pathways face significant energy barriers, advanced metabolic engineering is needed. For instance, redirecting ATP toward RubisCO carboxylation in E. coli has been reported to increase CO₂ fixation by 870%.³⁹ The cyanobacteria Synechocystis sp. PCC 6803′s capacity to fix carbon was increased by 3.2 times when the NAD(P) transhydrogenase gene was overexpressed to improve the reduced power supply.⁴⁰ In cell-free systems, ATP regeneration can be improved via engineered ATP synthases or light-driven proton pumps.⁴¹ Bio-electrochemical systems combining electrode-derived energy with microbial metabolism have also shown promise.⁴² However, excess ATP or NADPH can lead to metabolic stress and hinder growth, which may be resolved by combining this strategy with other techniques such as optimization of cell culture conditions.

3.1.2 Engineering of regulatory genes for CO₂ assimilation. Engineering of assimilation genes enhances CO₂ fixation efficiency in both natural autotrophs and engineered microbial systems. In the CBB cycle of cyanobacteria, the CO₂ fixation during day and night can be improved by deleting the regulatory protein CP12, which normally inhibits the activities of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and phosphoribulokinase (PRK) in the dark. Removing CP12 lifts this inhibition, allowing continuous enzyme activity and promoting more consistent carbon fixation from Ru5P to R15P across light and dark cycles.⁴³

In another study involving the rGly pathway in E. coli for enhanced growth on formic acid and CO₂, deletion of the ppsR gene encoding the regulatory protein of phosphoenolpyruvate synthase was implemented to block the conversion of pyruvate to phosphoenolpyruvate (PEP). This genetic modification prevented carbon loss through the reverse reaction and redirected carbon flux toward biomass formation, resulting in improved carbon assimilation and growth efficiency.⁴⁴

RubisCO expression in many autotrophs is directly controlled by the LysR-type transcriptional activator CbbR; elevating CbbR levels boosts transcription of the cbb operon and drives higher CO₂-fixation rates. In C. necator H16, an additional tier of control is provided by the RegB/RegA two-component system, which senses the cellular redox state and further modulates CBB cycle gene expression, ensuring that RubisCO and associated enzymes are expressed in tune with metabolic demands.⁴⁵ Artificial intelligence (AI)-driven omics analyses are increasingly important for identifying regulatory genes involved in CO₂ fixation. By analyzing large-scale genomic, transcriptomic, and metabolomic data, artificial intelligence helps uncover key regulatory elements. Manipulating these genes offers new opportunities to improve CO₂ assimilation in both natural and engineered microbial systems.

3.1.3 Enzyme engineering. Enzyme engineering approaches have emerged as transformative strategies to overcome the intrinsic limitations of natural CO₂ fixing enzymes and to boost carbon assimilation efficiency in both autotrophic organisms and engineered microbial platforms. By applying techniques such as site-directed mutagenesis, directed evolution, and rational design, researchers have been able to enhance the catalytic turnover and substrate specificity of key enzymes like RubisCO, whose sluggish kinetics and oxygenase activity limit overall carbon fixation rates.⁶ Directed evolution of RubisCO from Synechococcus PCC6301 enhanced its carboxylation activity by up to 85%. Similarly, heterologous expression of RubisCO from Thermosynechococcus elongatus BP1 in E. coli resulted in an improved carboxylation rate, efficiency, and CO₂/O₂ specificity. Expression of Rhodobacter sphaeroides RubisCO in E. coli resulted in a 27% increase in carboxylation rate and a 17% improvement in carboxylation efficiency, while maintaining stable CO₂/O₂ specificity; however, the holoenzyme biogenesis capacity was reduced by 40%. Furthermore, site-directed mutagenesis of R. sphaeroides RubisCO (C329A and A332 V) expressed in E. coli led to a 60% increase in carboxylation rate, a 22% improvement in carboxylation efficiency in air, and a 7% enhancement in CO₂/O₂ specificity.⁶ Beyond natural RubisCO variants, a synthetic CO₂-fixing enzyme, glycolyl-CoA carboxylase (GCC), was generated through rational design coupled with high-throughput microfluidic screening and microplate-based assays. GCC's catalytic efficiency improved by three orders of magnitude, achieving levels comparable to those of natural CO₂-fixing enzymes.⁴⁶ Taken together, these studies highlight both natural and synthetic strategies for enhancing CO₂ fixation efficiency beyond the limits of native RubisCO.

3.2 Process regulation of high cell-density culture

High cell density cultivation is critical in bioprocessing to boost the volumetric productivity of target products. High-density cultures face limitations from suboptimal carbon sources, substrate depletion, and cofactor shortages, which can be improved to increase growth and productivity.⁴⁷ High concentrations of substrates, such as glucose or sucrose, are often necessary to support biomass and product formation.⁴⁸

3.2.1 Optimization of carbon sources. The choice of carbon sources plays a crucial role in microbial growth and CO₂ fixation efficiency. Heterotrophic organisms like E. coli and Saccharomyces cerevisiae often require organic co-substrates to provide reduced power for CO₂ assimilation. For instance, a CBB-engineered E. coli strain showed a 45% increase in CO₂ fixation and improved biomass when grown on xylose instead of glucose. This strain was engineered with deletion of genes zwf and pfkAB which direct the flux towards the pentose phosphate pathway and glycolysis respectively. This strategy ensured that the flux shifted to favor the CBB pathway for carbon assimilation in a xylose medium.⁴⁹ Formate, a low-energy carbon source, was also optimized in autotrophic algal–bacterial consortia, leading to 0.15 g L⁻¹ biomass accumulation and 90% carbon assimilation efficiency under low light. Utilization of formate bypasses the gas–liquid mass transfer limitation of using bicarbonate as a carbon source due to its increased solubility and stability.⁵⁰ Although ideally, microbes would use CO₂ as the sole carbon source, its low solubility particularly as bicarbonate limits its uptake. This can be improved by delivering CO₂via pressurized gas or continuous gas streams, enhancing its bioavailability.⁵¹ However, such methods can alter culture pH and require careful monitoring.

3.2.2 Optimization of electron donors. Electron donors are vital for supplying the reducing power and energy required in microbial CO₂ fixation.⁵² Increasing their availability in the culture medium can enhance carbon capture. For instance, supplying high levels of hydrogen to Clostridium autoethanogenum improves CO₂ fixation by boosting the reduction potential.⁵³ While hydrogen-oxidizing bacteria efficiently fix CO₂ using H₂, the scalability is limited due to H₂ cost and safety concerns. Alternatives like formate or methanol show promise; for example, a Pichia pastoris strain engineered with the CBB cycle achieved autotrophic growth using methanol-derived reducing power, theoretically fixing up to 62 kg of CO₂.⁵⁴ Another strategy can be coupling microbes with electrochemical/photochemical systems which offers direct electron delivery, bypassing the need for organic donors.⁴² In one study, C. necator expressing rhodopsin was exposed to a light-driven photoelectric system and showed a 20% increase in CO₂ assimilation, replacing organic donors with an electrode for efficient electron transfer.⁵⁵ This photoelectric system replaces the need for organic electron donors with an electrode and drives maximum electron transfer efficiency in the strain. Inorganic donors like H₂, sulfide (S²⁻), and thiosulfate (S₂O₃²⁻) are also viable in heterotrophic systems. Combining multiple donors can create synergistic effects; for example, deep-sea chemoautotrophs achieved CO₂ fixation rates of 6.209 mg (L h)⁻¹ using a mix of NaNO₂, Na₂S₂O₃, and Na₂S.⁵⁶ Additionally, NCRCs such as NMN⁺ and NCD⁺ can substitute natural cofactors like NAD⁺ in engineered enzymes for efficient electron transfer.⁵⁷ Ultimately, optimizing energy flow, redox balance, and culture conditions is key to maximizing microbial CO₂ fixation, but strategies must be tailored to specific strains and process needs.

3.3 CRISPR-based genome editing in autotrophic microbes

CRISPR-based genome editing has emerged as a powerful tool to precisely modify the genomes of autotrophic microbes, thereby accelerating the development of strains with enhanced CO₂ fixation capabilities and improved bioproduct yields. In cyanobacteria such as Synechocystis sp. PCC 6803 and Synechococcus elongatus as well as in chemolithoautotrophs like C. necator, CRISPR-Cas systems have enabled marker-less gene knockouts, precise insertions, and transcriptional regulation via CRISPR interference (CRISPRi).^15,58–60 These advancements allow researchers to systematically dissect and rewire metabolic pathways, fine-tuning the expression of key genes involved in carbon assimilation and energy conversion. Moreover, the capacity for multiplex genome editing enables simultaneous modifications at several loci, which is critical for overcoming complex regulatory networks inherent to autotrophs. Despite challenges such as efficient DNA delivery and off-target effects, continuous improvements in CRISPR technology are rapidly expanding its utility in autotrophic systems, paving the way for the creation of robust microbial cell factories that can contribute to sustainable biofuel production and global carbon sequestration.

3.4 Adaptive laboratory evolution for improving CO₂ fixation efficiency

Adaptive laboratory evolution (ALE) has emerged as a powerful tool for improving CO₂ fixation efficiency and bioproduction yields across diverse microbial systems. ALE improved C. necator growth on volatile fatty acids (VFAs), with key mutations identified in phaR, the histidine kinase-response regulatory pair (H16_A1372/H16_A1373), and the tripartite transporter system (H16_A2296-A2298). These mutations enhanced growth, reduced NADH levels, and increased tolerance to higher VFA concentrations.⁶¹ Engineered E. coli strains, previously incapable of autotrophic growth, have 86 mutations and achieve CO₂ fixation with doubling times reduced to approximately 3.8 hours. Mutations in AtoA altered its substrate preference, enabling more efficient activation of acetoacetate with formyl-CoA, supporting better integration into the engineered pathway.³² Adaptive evolution increased the CO₂-specific growth rate from 0.008 h⁻¹ to 0.018 h⁻¹. Mutations were identified in the phosphoribulokinase (PRK) gene (A2G, T5A), which may influence ATP consumption and the supply of D-ribulose-5-phosphate for biosynthesis. Another mutation in nicotinic acid mononucleotide adenylyltransferase (NMA1; T358I) could affect intracellular NADH and ATP levels.⁵⁴

3.5 Omics-driven strategies (genomics, transcriptomics, and metabolomics)

Genomic analyses of model lithoautotrophs routinely confirm the presence of canonical CO₂ fixation operons. For example, the C. necator H16 genome encodes complete CBB cycle operons on both its chromosome and plasmid (including rbcL/S, prk, etc.). RNA-seq studies (DESeq2) under autotrophic (H₂/CO₂) versus heterotrophic conditions show hundreds of differentially expressed genes: overexpression of CBB regulators (CbbR/RegA) drives >2-fold induction of cbbL/S and adjacent CBB enzymes (with many TCA and PHB-biosynthesis genes also upregulated).⁶² The corresponding proteomic profiling by LC-MS/MS (e.g. MaxQuant) confirms these patterns: abundant RubisCO and other CBB enzymes are detected even under heterotrophic growth.⁶³ Targeted metabolomics (GC/LC-MS) in engineered cyanobacteria revealed altered pools of Calvin-cycle intermediates (for instance, elevated G3P and F15P with depleted 3-PG consistent with enhanced carbon fixation) and identified bottlenecks that were relieved by overexpression of GAPDH.⁶⁴ Finally, quantitative 13C–metabolic flux analysis mapped carbon flows through these pathways: e.g. steady-state 13C labeling in C. necator showed that the CBB cycle remained active (mixotrophically) even on organic substrates, with the Entner–Doudoroff pathway as the main sugar catabolism route and elevated oxidative pentose-phosphate flux feeding the CBB cycle.⁶⁵ Taken together, these multi-omics studies (genomics, transcriptomics, proteomics, metabolomics and fluxomics) quantitatively validate microbial CO₂ fixation machinery and pinpoint metabolic control points.

4. Synthetic biology application to non-traditional microbes and co-substrates for enhanced CO₂ fixation

4.1 Acetogens and sulfate-reducing bacteria

Acetogens, such as Acetobacterium woodii, Moorella thermoacetica, and C. autoethanogenum, and sulfate-reducing bacteria like Desulfovibrio desulfuricans and Desulfovibrio vulgaris, are anaerobic microorganisms capable of fixing CO₂via pathways such as the WLP and the rGly pathway. Metabolic engineering has significantly enhanced their CO₂ fixation efficiency and production of valuable products. For instance, ethanol production in C. ljungdahlii increased by 1.5-fold following adhE1 gene induction, while C. autoethanogenum saw a 1.2-fold increase in ethanol production with CODH/ACS gene overexpression.^66–68A. woodii doubled its ethanol production after adh4 gene deletion.⁶⁹D. desulfuricans fixed CO₂ through the reductive glycine pathway, the seventh known CO₂ fixation route. In this pathway, CO₂ was sequentially reduced to formate, then to glycine, which was further processed into acetyl-CoA via glycine reductase. Acetyl-CoA was then combined with another CO₂ to produce pyruvate.⁷⁰ Additionally, introducing the mevalonate pathway into C. ljungdahlii enabled isoprenoid production.⁷¹ These advancements highlight the potential of metabolic engineering in enhancing CO₂ fixation and biofuel production in acetogens and sulfate-reducing bacteria, with applications in bioenergy production and environmental remediation.

4.2 Methanotrophs and methylotrophs

Type II methanotrophs and certain methylotrophs utilize the serine cycle and the reductive glycine pathway for CO₂ fixation and energy metabolism, allowing efficient assimilation of C1 compounds. The yeast P. pastoris can grow on mixed C1 substrates, such as CO₂ combined with methanol or formate, using its native enzymes. This is achieved without the need for overexpression by redirecting methylene-THF flux toward glycine formation, accomplished through deletion of the mitochondrial serine hydroxymethyltransferase gene.¹¹ Type I methanotrophs can fix CO₂ using the CBB cycle. Methylomirabilis oxyfera encodes and actively transcribes genes of the CBB cycle, enabling carbon assimilation via this autotrophic pathway.⁷² CO₂ was essential for the growth of the RubisCO-encoding Methylococcus capsulatus Bath in a bioreactor with continuous gas inflow and outflow. RNA sequencing revealed active transcription of several carboxylating enzymes, including pyruvate carboxylase, phosphoenolpyruvate carboxylase, and ferredoxin oxidoreductase. Mutagenesis enabled the strain to assimilate CO₂ alongside methane, facilitating its growth.⁷³

5. Biotechnological applications and industrial potential

5.1 CO₂-based biomanufacturing of biofuels, bioplastics, and high-value chemicals

Microbial CO₂ sequestration technologies not only reduce carbon footprints but also serve as biorefineries, producing high-value chemicals such as bioethanol, polyhydroxyalkanoates (PHA), and succinate (Table 2). Microalgae and cyanobacteria fix CO₂via photosynthesis into biomass, which can subsequently be converted into biofuels like bioethanol, biodiesel, and biohydrogen.⁷⁴ Genetic engineering further enhances these processes; for example, S. elongatus has been modified to direct carbon flux toward fermentation pathways, increasing PHA production, while A. woodii achieves PHB accumulation up to 33.3% of cell content in high-pressure reactors.^75,76 Other acetogens facilitate chain elongation to generate medium-chain fatty acids, and engineered R. sphaeroides produces β-farnesene from CO₂ at levels 23 times higher than previous benchmarks. These examples highlight the potential of combining genetic technologies with biomass conversion as a strategy to mitigate CO₂ emissions while generating valuable chemicals.^77,78 Industrial implementation requires balancing microbial traits such as temperature tolerance, biomass yield, and carbon capture efficiency.⁷⁴

Table 2 Biomanufacturing valuable compounds using CO₂ as a carbon source

Product	Host strain	Metabolic engineering strategy	Cultivation conditions	Yield	Ref.
Ethanol	Saccharomyces cerevisiae	C. necator H16 was the source of the overexpressed genes cbbL1, cbbS1, and cfxP1, whereas S. cerevisiae D452–2 was the source of tHSP60 and tHSP10.	The yeast strains were inoculated into YP medium with 40 g L^−1 xylose at 0.5 g DCW L^−1 in 125 mL nitrogen-flushed, sealed serum bottles after being aerobically cultivated in YP medium and glucose, 10 ml and 20 g L^−1 respectively, for 24 hours at 30 °C. Anaerobic fermentation was subsequently conducted at 30 °C, 130 rpm.	14.5 g L^−1	79
1,2-Propanediol	Synechocystis sp. PCC6803	To convert dihydroxyacetone phosphate to methylglyoxal, which is subsequently reduced to acetol and then to 1,2-propanediol, native genes adh (alcohol dehydrogenase), yqhD (aldehyde reductase), and mgsA (methylglyoxal synthase) were overexpressed.	Synechocystis was grown on BG11 or YBG11 agar with antibiotics under 25 μmol m^−2 s^−1 light, ambient CO2, 30 °C, and 75% humidity for 10 days. Then it was transferred to YBG11 with 50 mM HEPES; liquid cultures (50 mL) were grown in 250 mL baffled flasks at 30 °C, 150 rpm, 2% CO2, increasing light, and 75% humidity; main cultures were inoculated at OD750 = 0.2.	1 g L^−1	80
Ethylene	Synechocystis sp. PCC6803	Overexpression of phosphoenol pyruvate carboxylase.	For two to three days, Synechocystis strains that overexpressed native gene or carried a kanamycin resistance cassette were cultivated in BG11 medium with kanamycin.	16.77 ± 4.48 μg per mL per OD per day	81
PHB	Synechococcus elongatus UAM-C/S03	N/A	Nitrogen limiting conditions; the biomass was washed to remove nitrogen and resuspended in 2 × BG11; experiments at pH ∼8 or uncontrolled pH were conducted at 100 μmol photons m^−2 s^−1, 35 ± 0.5 °C, with 0.5 g L^−1 initial biomass; batch cultures lasted 7–9 days with daily sterile sampling.	40.82 mg L^−1 d^−1	82
PHA	Cupriavidus necator H16	To produce P(3HB), a recombinant R. eutropha PHB-4 strain was created, carrying the plasmids pBBR1:phaCABRe or pBBR1-phaRCYB4AB with PHA synthase genes from R. eutropha and Bacillus cereus YB-4, respectively, under the control of PHA promoter.	A 250 mL jar fermenter with continuous gas flow was used for autotrophic culture in NR medium. Using mass flow controls, a gas mix (H2 : O2 : CO2 : N2 = 3.8 : 7.3 : 13.0 : 75.9) that was created by combining 5.8% H2, 19.9% CO2, and air was delivered at 1–15 mL min^−1, keeping H2 below explosive values.	2.94 g L^−1	83
Acetic acid	Clostridium acetobutylicum	Carbon monoxide dehydrogenase was overexpressed.	A 0.6 V poised potential at the working electrode was used to operate a small-scale microbial electrochemical system (100 mL total, 75 mL working volume); strains were grown in the appropriate media and inoculated at OD 0.6 (10 v/v %), with CO2 supplied in the headspace without disturbing biofilms; the retention time was 60 h for CO2 fixation and conversion analysis.	7.8 g L^−1	84
Reservatrol	Cupriavidus necator H16	Tyrosine ammonia lyase, 4-coumaroyl CoA ligase, and stilbene synthase are heterologous genes for resveratrol production that were introduced into C. necator H16. Multiple copies of Vitis vinifera's stilbene synthase, interruption of the PHB pathway, and overexpression of acetyl-CoA carboxylase	C. necator H16 was routinely grown in LB at 30 °C and 200 rpm. To produce heterotrophic resveratrol, strains were cultivated in 40 mL of minimal medium (MM) with 10 g L^−1 fructose and 5 mM tyrosine. For lithoautotrophic fermentation, precultures in MM with fructose were washed and inoculated into MM with tyrosine in 157 mL serum bottles (20 mL volume), pressurized daily with H2 : O2 : CO2 gas (70 : 20 : 10 or 78 : 2 : 10, 150 kPa), agitated at 200 rpm and 30 °C with original OD = 1 or 5. After 24 hours, gene expression was induced with 0.2% L-arabinose, and kanamycin (200 μg mL^−1) was added for plasmid maintenance.	1.9 mg L^−1	85
Extracellular polysaccharide (EPS)	Porphyridium purpureum CoE1	N/A	A 10% inoculum size was used to cultivate P. purpureum CoE1 in a flask of 1 L capacity with 500 mL of artificial seawater (ASW). For 18 days, the flask was kept in the same illumination incubator and exposed to cool white, fluorescent light sources that emitted about 165 μmol photons m^−2 s^−1. A ventilation rate of 1 L min^−1 was used to maintain the culture temperature at 25 °C. Different dosages of inorganic salt were used to create ASW. Throughout the cultivation process (18 days), the dosage was fixed at 30%, 50%, or 70%.	2.89 g L^−1	86
Oxalate	Escherichia coli	Genes encoding fumarate and nitrate reductase, lactate dehydrogenase, pyruvate carboxylase, oxaloacetate acetylhydrolase, and acetyl-CoA synthetase were expressed heterologously	E. coli BL21 (DE3) was grown in LB medium aerobically and in TB medium anaerobically.	23 μM	87
Acetone	Clostridium autoethanogenum	Overexpression of secondary alcohol dehydrogenase (sAdh) variants and -CoA transferase (CtfAB)	In 2 L CSTR reactors, continuous fermentation was carried out anaerobically at 37 °C using a condenser to reduce volatile product loss and a synthetic gas blend (50% CO, 10% H2, 30% CO2, and 10% N2) at atmospheric pressure. Pilot tests using a mixture of 50% carbon monoxide, 10% hydrogen, 20% carbon dioxide, and 20% nitrogen gases were carried out in a bespoke reactor at 1.4 bar. For gas flow rates of 200–800 SCCM for the CSTRs and 11.5–23 NLPM for the pilot reactors, the medium dilution rates varied from 1.0 to 6.5 d^−1.	3 g L^−1 h^−1	88
Lactic acid	Pichia pastoris	Integration of one CYB2 knockout, which codes for L-lactate cytochrome c oxidoreductase, and a bacterial lactate dehydrogenase gene	Using 0.5% methanol and 5% CO2 in the intake gas, bioreactors were inoculated from an YPG preculture to an OD of 2 for autotrophic cultivation. Following the initial sample, the methanol content was raised to 1%.	600 mg L^−1	89
Succinate	Synechococcus elongatus PCC 7942	Blocking glycogen synthase and succinate dehydrogenase using CRISPRi	S. elongatus strains were grown in 60 mL of BG-11 medium containing IPTG in glass tubes for production with ambient or CO2-enriched air, beginning at OD730 ≈ 0.2. After being grown to an OD730 of 0.4–0.6, the pre-culture was stimulated with IPTG, centrifuged to concentrate it, and then resuspended. The temperature was set at 30 °C, the intensity of light was kept at 200 μE s^−1 m^−2, and 100 mL min^−1 CO2-enriched air was provided. BG-11 medium was replaced and the pH was kept at 10 with NaOH.	8.9 g L^−1	90

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5.2 Integration with industrial waste streams and direct air capture (DAC) systems

A key advantage of microbial CO₂ mitigation is its compatibility with industrial waste gases and direct air capture (DAC) systems. Industrial emissions from sectors like cement, steel, and power plants contain high CO₂ levels that can be directly used by microbial cultures. For instance, engineered S. elongatus has been used to convert flue gas into bioethanol.⁹¹ Integrating such systems with emission treatments can reduce waste and generate valuable products. DAC, although energy-intensive,⁹² becomes more sustainable and cost-effective when paired with microbial fixation.⁹³ Microalgae like Chlorella vulgaris, Spirulina sp., and Botryococcus braunii have shown carbon fixation rates of 50 g m⁻² day⁻¹ using DAC-derived CO₂.⁹⁴ Additionally, a chemoenzymatic system has converted captured CO₂ into pyruvate and amino acids (glycine and serine) via the reductive glycine pathway, highlighting the potential of integrating biological CO₂ sequestration with both industrial emissions and DAC.⁹⁵

5.3 CO₂ mitigation and biomass conversion

To reduce reliance on fossil resources, lignocellulosic biomass has emerged as a promising renewable carbon source for the sustainable production of chemicals and materials. Microorganisms play a central role in this transition by converting residual biomass and CO₂ into fuels and value-added products through natural and engineered pathways. In industrial practice, microbial processes that transform lignocellulose into biogas mirror those naturally occurring in the environment. Heterotrophic bacteria and fungi degrade lignocellulosic and protein-rich residues into biogas (CH₄ + CO₂), organic acids, and nutrient-rich compost, while methanogens and acetogens re-capture part of the released CO₂ to generate additional methane or acetate, thereby enhancing carbon recovery.^96,97

Photosynthetic microbes such as cyanobacteria and microalgae further contribute by fixing CO₂ into cellular biomass that can be harvested for biofuels, animal feed, or fertilizers. Advances in gene editing and synthetic biology have extended their utility to the sustainable biosynthesis of high-value compounds, including olefins, cinnamaldehyde, and curcumin.⁹⁸ At the process scale, a 10–20 L bubble column bioreactor using Bacillus cereus SSLMC2 achieved biomass yields of 9.14–10.78 g L⁻¹ and CO₂ removal efficiencies of 56–85%. FT-IR and GC-MS analyses identified value-added products such as carboxylic acids, fatty alcohols, and hydrocarbons, while techno-economic assessment demonstrated low-cost operation ($0.167–0.187 per g C treated), underscoring the feasibility of CO₂ bio-mitigation strategies.⁹⁹ Integrating these microbial platforms with biomass valorization and CO₂ capture provides a cornerstone for the circular bioeconomy, enabling the conversion of residual biomass and anthropogenic CO₂ into renewable energy carriers and bio-based materials. Such integration offers a practical route to reduce net greenhouse gas emissions and advance toward carbon neutrality.

5.4 Challenges and future directions in industrial scalability

Microbial CO₂ fixation faces persistent challenges, largely stemming from enzymatic limitations that make biological carbon assimilation slower than chemical catalysis. Low CO₂ solubility and dissolution rates further restrict its bioavailability and uptake.⁷⁷ Natural phototrophs demand high levels of reducing power in the form of NADPH and ATP, and their cultivation requires photobioreactors and tightly controlled environments, resulting in elevated operational costs.¹⁰⁰ Engineered heterotrophic strains offer greater pathway flexibility, but their performance often declines over time due to genetic instability and metabolic imbalances. In addition, heterotrophs typically prioritize reduced carbon substrates over CO₂, which diminishes fixation efficiency.¹⁰¹ Thermodynamic constraints inherent to pathways such as the WLP and rTCA cycle further complicate the optimization of carbon flux toward biomass or product formation.¹⁰²

Selecting an effective CO₂-fixation route requires weighing several interdependent trade-offs. One central consideration is the choice of host organism. Native autotrophs such as cyanobacteria, acetogens, and hydrogen-oxidizing bacteria provide proven CO₂ assimilation pathways and exhibit physiological robustness, whereas synthetic hosts including engineered E. coli or yeast offer broader flexibility for pathway engineering and product diversification, but often demand extensive optimization to achieve stable performance.¹⁰³ Another key factor concerns the balance between productivity and carbon accounting. Autotrophic processes maximize the fraction of carbon incorporated directly from CO₂, strengthening their sustainability profile, yet they generally operate at slower production rates and lower overall efficiency. In comparison, mixotrophic approaches that combine CO₂ with energy-rich organic substrates can achieve higher titers and productivities, although this complicates carbon balances and introduces trade-offs in life-cycle assessments.³⁶ Finally, the mode of redox supply defines the operational boundaries of the process. Electrobiological platforms couple microbial metabolism directly to electrodes, either through direct electron uptake or redox mediators, allowing precise control over reducing power delivery. Gas-fed systems, in contrast, provide chemical reductants such as H₂ (and in some cases CO or CH₄), which can facilitate reactor integration at scale but impose additional challenges associated with gas liquid mass transfer limitations and safety considerations.¹⁰⁴ Collectively, these considerations indicate that there is no universal optimum for biological CO₂ fixation. Instead, the choice of strategy should be aligned with both the desired product profile and the industrial context (Table 3). For instance, formatotrophic or rGly pathway based E. coli strains may be better suited for flexible chemical production platforms, whereas hydrogenotrophic chassis are more appropriate for high-yield protein or single-cell protein applications.^25,105 Biopolymer production relies on polymer-accumulating microbes like C. necator with metabolic flux redirected toward polymer pathways.

Table 3 Comparative overview of natural, synthetic, electrobiological, and cell-free CO₂-fixation strategies

Representative organisms	Carbon & energy source(s)	Doubling time	Typical product yields or titers	Key strengths	Key limitations	Scale maturity	Ref.
Natural microbial systems
Clostridium autoethanogenum, Clostridium ljungdahlii	CO2/CO + H2 or syngas	∼6–8 h under optimized conditions	Ethanol titers >40 g L^−1 in industrial plants	Commercially proven gas fermentation, high carbon incorporation, tolerance to CO	Gas–liquid mass transfer limitations, strict anaerobiosis, limited genetic tools	Commercial deployment for ethanol (LanzaTech)	107
Cupriavidus necator	CO2 + H2 + O2	∼2–4 h	Protein yields ∼65–70% of dry weight, SCP productivity up to 20 g L^−1 DCW	High protein content, scalable SCP production, aerobic operation	Requires H2/O2 mixing, safety hazards, limited product diversity	Pilot scale demonstration	108
Cyanobacteria (Synechocystis sp. PCC 6803), microalgae	CO2 + light	6–12 h	PHB up to 20–30% DCW; biomass productivity ∼0.5–1 g L^−1 day^−1	Renewable inputs, long research history, genetic tractability in models	Light distribution limits, low volumetric productivity, require large reactor footprint	Long research history, niche applications, no large-scale commodity production	109
Synthetic microbial systems
Engineered E. coli with the reductive glycine pathway	CO2 + formate (electrosynthesized)	<4.5 h	3.8 g L^−1 mevalonate, demonstration of isoprenol and terpenoids	Broad metabolic engineering scope, oxygen tolerance, rapid ALE improvements	Relies on electrochemical formate efficiency, pathway stability, scale not yet proven	Lab-scale demonstrations	25 and 110
Electrobiological systems (MES)
Sporomusa ovata, acetogenic consortia	CO2 + electrons from electrodes (direct or mediated)	Reported >12 h (biofilm-limited)	Acetate titers 10–20 g L^−1 in lab-scale systems	Direct coupling to renewable electricity, precise redox control, avoid H2 handling	Low areal productivity, electrode–biofilm interface resistance, high materials cost	Lab- to bench-scale; early pilot tests	111
Cell-free synthetic cycles
CETCH cycle, THETA cycle, self-replenishing modules	CO2 + NAD(P)H/ATP (regenerated chemically/electrochemically)	Not applicable (enzyme systems)	The CETCH cycle achieves CO2 fixation rates up to 5× faster than RubisCO; turnover ∼5 mmol CO2 g^−1 enzyme h^−1	High specific activity, no biomass maintenance, modular design	Enzyme stability and cost, continuous cofactor regeneration, integration challenges	Laboratory scale proof-of-concept	46 and 112

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Recent advances aim to overcome current barriers and improve industrial feasibility. ALE and continuous culture optimization are used to enhance strain robustness and long-term stability.¹⁰⁶ Synthetic biology enables the development of more efficient carboxylases, including engineered RubisCO variants and artificial CO₂-fixing enzymes, to accelerate assimilation. Electro-biotechnology is emerging as a promising route that couples microbial metabolism with renewable electricity to drive CO₂ conversion. To alleviate solubility constraints, new delivery systems such as membrane-based gas transfer are under development. At process scale, cost-effective reactor designs and integration with carbon capture technologies are essential to repurpose industrial CO₂ emissions into fuels, polymers, and biomaterials.

When evaluated through the framework of Green Chemistry, CO₂-fixation strategies show distinct alignments with core principles. Table 3 integrates natural, synthetic, electrobiological, and cell-free systems to highlight technological progress and sustainability trade-offs. Gas-fed microbial systems achieve high atom economy but rely on low-carbon H₂ or CO inputs and efficient gas transfer to maintain performance. Synthetic formatotrophy enables transparent carbon accounting and flexible pathway design, although its overall efficiency is constrained by the electricity-to-formate conversion step. Electrobiological platforms eliminate the need for molecular reductants and couple directly to renewable electricity, yet they remain limited by technological immaturity and scale. Cell-free cycles offer the prospect of superior atom economy and precise flux control, provided challenges in enzyme stability and cofactor regeneration can be addressed. Phototrophic systems operate entirely on renewable inputs, but their productivity is restricted by light distribution and reactor design. Future work should therefore prioritize comparative benchmarking of carbon yields, energy requirements, and life cycle impacts to ensure that advances in microbial CO₂ fixation translate into verifiable environmental benefits consistent with the principles of Green Chemistry.

6. Conclusion

Microbial CO₂ fixation has matured from reliance on native autotrophic cycles such as the CBB, reductive TCA, Wood–Ljungdahl, and archaeal HP/HB pathways to embrace engineered enhancements, wholly synthetic routes (e.g., CETCH, formolase, and SACA), electro-microbial hybrids, and non-traditional hosts like acetogens, methanotrophs, methylotrophs, and designer yeasts, forging a versatile toolkit for carbon capture and bioproduction; yet, translating these innovations into robust, industrially viable processes demands overcoming interrelated hurdles, namely, boosting enzyme turnover and thermodynamic favorability to elevate pathway flux, securing balanced ATP and redox cofactor supply through novel co-substrate or chemical strategies, expanding reliable genetic toolboxes and dynamic regulatory circuits for strict anaerobes and extremophiles, mitigating gas liquid mass transfer bottlenecks and genetic instability in high-density bioreactors, integrating predictive multi-omics and reactor modeling (potentially via AI-driven digital twins) to streamline design–build–test cycles, and anchoring development in rigorous techno-economic and life-cycle analyses; only through coordinated advances in enzyme and pathway engineering, systems biology, process intensification, computational modeling, and supportive policy frameworks can we realize scalable, cost-effective, and sustainable CO₂ to chemical biorefineries. Looking forward, two research priorities stand out: (i) engineering CO₂-fixing enzymes and synthetic pathways with higher catalytic efficiency and (ii) developing integrated reactor platforms that couple advanced control, multi-omics, and techno-economic assessment to ensure scalability. Guided by these directions and supported by advances in systems biology, process intensification, and policy, microbial CO₂ fixation can evolve into cost-effective and sustainable CO₂-to-chemical biorefineries.

Author contributions

Zeeshan Mustafa: conceptualization and writing – original draft. Naeem Auroona: conceptualization and writing – original draft. Arslan Sarwar: writing – original draft. Eun Yeol Lee: conceptualization, writing – review & editing, supervision, project administration, and funding acquisition.

Conflicts of interest

The authors declare no conflicts of interest.

Data availability

No primary research results, software or code have been included and no new data were generated or analysed as part of this review.

Acknowledgements

This work was supported by the National Research Foundation of Korea (NRF) grant funded by the Korean government (MSIT) (RS-2024-00466473). This research was supported by the Bio&Medical Technology Development Program of the National Research Foundation (NRF) funded by the Korean government (MSIT) (No. RS-2025-02214232).

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Footnote

† These authors contributed equally to this work.

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