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Biochemistry
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Biochemistry
Blended nexus molecules promote CO2 to l-tyrosine conversion
l-tyrosine production from CO2
https://orcid.org/0000-0002-2509-6198
Fan Lei Conceptualization Formal analysis Investigation Methodology Resources Validation Visualization Writing - original draft Writing - review & editing 1 †
https://orcid.org/0009-0005-7998-1642
Zhu Zihan Conceptualization Data curation Formal analysis Investigation Methodology Resources Validation Visualization Writing - original draft Writing - review & editing 2 †
https://orcid.org/0000-0002-5282-3194
Zhao Siyan Formal analysis Methodology Project administration 2
https://orcid.org/0000-0003-4052-3335
Panda Smaranika Conceptualization Investigation Methodology Project administration Resources Validation Visualization Writing - review & editing 1
https://orcid.org/0000-0002-7527-9056
Zhao Yilin Formal analysis Methodology Validation Visualization Writing - original draft Writing - review & editing 1
Chen Jingyi Conceptualization Visualization Writing - original draft Writing - review & editing 1
Chen Lei Conceptualization Formal analysis Investigation Methodology Validation Visualization 1
Chen Junmei Methodology Validation 1
https://orcid.org/0000-0003-1895-6628
He Jianzhong Conceptualization Data curation Funding acquisition Methodology Project administration Resources Supervision Validation Visualization Writing - review & editing 2
https://orcid.org/0000-0003-0606-7030
Zhou Kang Conceptualization Project administration 1
https://orcid.org/0000-0002-1931-7767
Wang Lei Conceptualization Data curation Formal analysis Funding acquisition Investigation Methodology Project administration Resources Supervision Validation Visualization Writing - original draft Writing - review & editing 1 *
1 Department of Chemical and Biomolecular Engineering, National University of Singapore, 4 Engineering Drive 4, Singapore 117585, Singapore.
2 Department of Civil and Environmental Engineering, National University of Singapore, 1 Engineering Drive 2, Singapore 117576, Singapore.
* Corresponding author. Email: wanglei8@nus.edu.sg
† These authors contributed equally to this work.

06 9 2024
06 9 2024
10 36 eado135218 1 2024
31 7 2024
Copyright © 2024 The Authors, some rights reserved; exclusive licensee American Association for the Advancement of Science. No claim to original U.S. Government Works. Distributed under a Creative Commons Attribution NonCommercial License 4.0 (CC BY-NC).
2024
The Authors
https://creativecommons.org/licenses/by-nc/4.0/ This is an open-access article distributed under the terms of the Creative Commons Attribution-NonCommercial license, which permits use, distribution, and reproduction in any medium, so long as the resultant use is not for commercial advantage and provided the original work is properly cited.

Using CO2 as the primary feedstock offers the potential for high-value utilization of CO2 while forging sustainable pathways for producing valuable natural products, such as l-tyrosine. Cascade catalysis is a promising approach but limited by stringent purity demands of nexus molecules. We developed an abiotic/biotic cascade catalysis using blended nexus molecules for l-tyrosine synthesis. Specifically, we begin by constructing a solid-state reactor to reduce CO2 electrochemically, yielding a mixture of acetic acid and ethanol, which serves as the blended nexus molecules. Subsequently, we use genetic engineering to introduce an ethanol utilization pathway and a tyrosine producing pathway to Escherichia coli to facilitate l-tyrosine production. The ethanol pathway synergistically cooperated with the acetic acid pathway, boosting l-tyrosine production rate (nearly five times higher compared to the strain without ethanol utilization pathway) and enhancing carbon efficiency. Our findings demonstrate that using blended nexus molecules could potentially offer a more favorable strategy for the cascade catalysis aimed at producing valuable natural products.

Blended nexus molecules enable enhanced CO2 to l-tyrosine conversion in biotic/abiotic cascade catalysis.

http://dx.doi.org/10.13039/501100001348 Agency for Science, Technology and Research U2102d2002 http://dx.doi.org/10.13039/501100001352 National University of Singapore A-0009176-02-00, A-0009176-03-00 http://dx.doi.org/10.13039/501100001352 National University of Singapore CHI-P2022-06 National Research Foundation (NRF) Singapore NRF-NRFF13-2021-0007, NRF-CRP27-2021-0004 Centre for Hydrogen Innovations CHI-P2022-06 License OptionCC BY-NC
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pmcINTRODUCTION

l-tyrosine, one of the 20 essential amino acids, is crucial for protein synthesis in all living organisms, and it has been extensively used in pharmaceuticals, food industry, feed industry, and dietary supplements (1, 2). For instance, l-tyrosine is a key precursor for synthesizing various high-value products, including the Parkinson’s disease drug 3,4-dihydroxy-l-phenylalanine, melanin, and a wide range of other bioactive compounds such as flavonoids, alkaloids, and stilbenoids (3). Thus, there is a growing interest in advancing technologies to achieve the sustainable production of l-tyrosine (4). In general, l-tyrosine could be produced through four methods, including protein hydrolysis, chemical synthesis, enzymatic conversion, and fermentation (5). Previously, l-tyrosine was extracted from protein hydrolysates. However, the energy-intensive separation and the subsequent purification processes often result in high energy cost and low product yields, making this approach inadequate to meet the rising demand of tyrosine (6). The chemical synthesis of l-tyrosine typically involves multiple steps including functional groups protection and deprotection, and this process leads to a racemic mixture, necessitating intricate chiral separation procedure (7). Enzymatic synthesis of l-tyrosine using tyrosine-phenol lyases offers the advantage of achieving high regio- and stereo-selectivity. However, the enzyme’s instability and sharp decline in activity after prolonged catalysis at high temperatures pose challenges for its practical application (5). In addition, the substrate pyruvate is conventionally generated through the dehydrative decarboxylation of tartaric acid at high temperatures, making the whole process high cost and environmentally unsustainable (7). On the other hand, the emergence of large-scale fermentation has established microbial production using genetically engineered strains, such as Escherichia coli, as a promising method for l-tyrosine production (8). However, one major limitation of this approach is that the feedstocks, such as sugar (i.e., glucose), are often valuable molecules, leading to high production cost (6, 9, 10). In addition, the biosynthetic pathways using these starting materials involve multiple metabolic branch nodes, which can result in drastic carbon loss, low product yield, and the formation of byproducts (11).

In nature, plants can convert atmospheric CO2 into aromatic amino acids, including l-tyrosine, phenylalanine, and tryptophan (4), through the process of photosynthesis driven by solar energy. Mimicking the natural process of converting CO2 into valuable products (i.e., amino acids) holds great potential to revolutionize the food and pharmaceutical industries, while offering a promising approach in mitigating the pressing climate challenges caused by the massive CO2 emission (12). Emerging technologies, such as electrochemical CO2 reduction (CO2R), have received extensive attention for their potential to use renewable electricity as energy sources to upgrade CO2 into valuable products (13–20). However, in CO2R, the resulting carbon chains in the products are often short, encompassing compounds like formate, carbon monoxide, ethanol, acetate, and n-propanol (21). It remains a grand challenge to convert CO2 into molecules larger than three carbons at high production rates due to the complicated reaction pathways (22). In contrast, microbial-assisted electrosynthesis, achieved by directly attaching the microbes to the photo/electro-cathode, has shown notable selectivity toward multicarbon products, including butyrate and isobutanol (23). However, a major challenge arises from the disparities between the biochemical and electrochemical systems, as the microbials can only withstand very low operating current densities (24). This limitation poses a drastic hurdle in achieving high production rates, which are essential for practical applications. Cascade process built on spatially decoupled electrochemical and biochemical reactors offers an opportunity to overcome this limitation. The feasibility of such cascade systems has been recently demonstrated by coupling an electrochemical CO2R reactor with a fermentation module, resulting in the production of various long-chain chemicals including butanol, hexanol, glucose, fatty acids, polyesters, sugar, and β-farnesene (25–31). However, the synthesis of amino acid from CO2 powered by renewable energy sources has not been demonstrated to the best of our knowledge, motivating further research on expanding the variety of product species (table S1). There is also a need to expand the nexus species (identity/purity) that connect electrocatalysis and biocatalysis to achieve greater flexibility in such systems and high carbon utilization efficiency. This is particularly important if the nexus molecules are obtained from CO2R, due to its unsatisfactory selectivity toward single products.

The key challenge associated with the spatially decoupled abiotic/biotic cascade catalytic system design mainly arises from the disparity between the electrolysis and biocatalysis systems, attributed to the diversity, concentration, and impurity effects of the nexus molecules (Fig. 1A). In addition, achieving the maximized utilization of nexus molecules by the microbes may require the incorporation of multiple biosynthetic pathways. Here, we present a cascade catalysis design that uses blended molecules as nexus molecules, different from recent works in cascade catalysis (25–31), where single nexus molecule was used in microbial fermentation. The blended nexus molecules, devoid of any other impurities, can be directly obtained from electrocatalysis through an optimized electrocatalytic system using solid electrolyte (Fig. 1B). These strategies eliminate the necessity for an additional product separation process before feeding the molecules directly into the biotic reactor. The blended molecules induce cooperative effects in the engineered strain, resulting in an obviously increase in l-tyrosine production. Specifically, we carried out genetic engineering on E. coli strains to incorporate an additional pathway for synthesizing l-tyrosine using ethanol as the feedstock, while preserving the existing acetate-based pathway. The incorporation of the ethanol utilization pathway stimulates the growth of the E. coli strains, and this, in turn, leads to a simultaneous enhancement of the acetate pathway for l-tyrosine production. Overall, our strategies, which are based on blended nexus molecules involving electrocatalysis engineering, the incorporation of multiple biosynthetic pathways, and integrated system optimization, present a promising avenue for the valorization of waste CO2 into valuable chemicals, such as l-tyrosine.

Fig. 1. Schematic illustration of the l-tyrosine synthesis from CO2 using blended nexus molecules based on an abiotic/biotic cascade catalytic system.

(A) The primary challenge in abiotic/biotic cascade catalysis stems from the disparity between the supply of electrocatalysis and the demand of nexus molecules for biotic catalysis. (B) Genetic engineering is used to address the above disparity. Electrocatalytic engineering, involving a solid electrolyte reactor and a recycled products flow, is used to prevent salt mixture and increase the products concentration. Specifically, CO2 is first converted to pure acetic acid and ethanol solution powered by renewable electricity through a two-step electrolysis, then the electrochemically produced blended nexus molecules containing acetic acid and ethanol are directly used by the genetically engineered E. coli in a biological reactor for the sustainable synthesis of l-tyrosine. The cooperation effects between ethanol utilization pathway and acetic acid utilization pathway in the engineered E. coli enhance the production of l-tyrosine.

RESULTS

Electrocatalytic engineering for improved production of blended nexus molecules

Electrochemically converting CO2 to CO with high selectivity and activity are relatively straightforward (figs. S1 and S2) and high-rate CO2R to CO has been developed using various catalysts (32, 33). Hence, in this study, we focus on the catalyst and reactor design toward electrochemical reduction of CO (COR) to acetate and ethanol, as improved selectivity of these products has been demonstrated in COR (34–37). Cu-based catalysts exhibit considerable selectivity toward multicarbon hydrocarbons (i.e., ethylene) in the COR; however, the selectivity and productivity toward oxygenates (acetate and ethanol) are often modest (34, 36, 38). Modifying Cu with other metals (i.e., Ag, Au, and Pd) to afford Cu-based bimetallic catalysts, presents an appealing strategy to tune the catalytic performance for CO2R/COR (16, 18, 37, 39–41). In particular, recent works show that Cu-Ag interfacial sites exhibit reduced affinities to C and OH, thus promoting the production of C2 oxygenates (35, 37, 42–45). Inspired by these findings, we prepared a range of CuAg nanoparticle catalysts with different Cu/Ag ratios aiming to achieve optimized production of acetate and ethanol from COR. The CuAg catalysts were synthesized through a procedure of coprecipitation followed by in situ electroreduction, designed for straightforward future large-scale production. Elemental mapping based on transmission electron microcopy (TEM) (figs. S3 to S7) confirms the presence of Cu and Ag elements within the nanoparticles. The apparent uniform distributions of the Cu and Ag indicate a substantial presence of the Cu/Ag bimetallic interface, which is anticipated to favor the oxygenate production. In addition, x-ray photoelectron spectroscopy (XPS) and powder x-ray diffraction (XRD) patterns of the CuAg catalysts also demonstrate the existence of metallic Cu and Ag (figs. S8 to S10).

We first assessed the COR performance of CuAg catalysts with different Cu/Ag ratio in a typical three-electrode flow reactor using 1.0 M potassium hydroxide (KOH) as the electrolyte. Among the different CuAg candidates, CuAg-2 (surface Cu-to-Ag atomic ratio of ~1.5 was determined by XPS; fig. S8) exhibited the highest C2 oxygenates Faradaic efficiencies (FEs) and partial current densities (Fig. 2A and figs. S11 to S16), likely due to the optimized Cu-Ag interfacial active sites. At high current density of 800 mA cm−2, CuAg-2 reached a remarkable FE exceeding 51% toward C2 oxygenates (Fig. 2A), including 30% FE toward acetate and 21% FE toward ethanol (fig. S13). At the same conditions, pristine Cu only exhibited 23% FEs toward C2 oxygenates (fig. S11). In addition, CuAg-2 exhibited C2 oxygenates partial current density 437 mA cm−2 at −1.1 V versus reversible hydrogen electrode (RHE), which was nearly twice as high as that of the pristine Cu nanoparticles (fig. S16).

Fig. 2. Electrochemical CO reduction performance of CuAg catalyst.

(A) FEs of C2 oxygenates of Cu-Ag catalysts with different atomic ratio at various current densities. (B) Schematic illustration of the solid-state electrochemical COR reactor. (C) FEs of different products at various current densities on CuAg-2 in the solid-state electrochemical COR reactor. (D) C2 oxygenates partial current densities at different cell voltage. (E) The production rates of C2 oxygenates at different current densities. (F) Stability test of the solid-state reactor at 300 mA cm−2.

CuAg-2 catalyst exhibited a noteworthy C2 oxygenates partial current density, demonstrating its potential to generate concentrated C2 oxygenates streams suitable for fermentation. However, the resultant acetate and ethanol were mixed with a highly concentrated salt electrolyte (1.0 M KOH), rendering the solution unsuitable for direct microbe growth (11, 27). To overcome this issue, we used a solid electrolyte electrolyzer recently reported for pure C2 oxygenates production (46–49). As depicted in Fig. 2B, electrochemically generated acetate/ethanol and protons are transported across the ion exchange membranes from the cathode and anode, respectively, into the porous solid electrolyte layer. The acetate is then protonated to form acetic acid. Subsequently, the generated C2 oxygenates are carried out by the deionized (DI) water stream. Within this configuration, maximum C2 oxygenates FEs of 50% was achieved at 600 mA cm−2 (Fig. 2C). Despite the plateau of FEs, the partial current density and production rate toward C2 oxygenates could be further boosted to 463 mA cm−2 and 220 mg hour−1 cm−2 (~3.3 mmol hour−1 cm−2), respectively (Fig. 2, D and E, and fig. S17). Last, we also assessed the stability of the system for continuously C2 oxygenates production. As shown in Fig. 2F, we demonstrate a continuous and stable COR electrolysis over 100 hours at 300 mA cm−2, negligible degradations in both cell voltage and C2 oxygenates product selectivity are observed.

Metabolic engineering and substrate optimization for enhanced fermentation

As discussed above, the production of acetic acid and ethanol via electrochemically CO2/CO reduction is attainable. However, wild strain of E. coli cannot spontaneously use both acetic acid and ethanol for l-tyrosine production, owing to the following reasons: first, the native E. coli is unable to grow using ethanol as the sole carbon and energy source, which is possibly attributed to the low transcription rate of AdhE (a protein involved in ethanol metabolism, encoded by the adhE gene) and its sensitivity to metal ion-catalyzed oxidation (50, 51); second, l-tyrosine production in native E. coli strains was tightly regulated (52). Although acetic acid and ethanol have been used individually as carbon sources for microbial fermentation (51, 53, 54), the utilization of both acetic acid and ethanol within a single strain has not been demonstrated (table S1). To overcome these challenges and enhance the efficiency of l-tyrosine production using blended nexus molecules, we developed an engineered E. coli strain by integrating the ethanol utilization pathway with the l-tyrosine-producing pathway (Fig. 3).

Fig. 3. Key metabolic pathway converting acetic acid and ethanol into l-tyrosine.

OAA, oxaloacetate; PEP, phosphoenolpyruvate; 2-PG, 2-phosphoglycerate; G3P, glyceraldehyde 3-phosphate; E4P, erythrose 4-phosphate; DHAP, 3-deoxy-d-arabino-heptulosonate 7-phosphate; HPP, 4-hydroxyphenylpyruvate; ATP, adenosine triphosphate. Adh2 encodes alcohol dehydrogenase. Ada encodes acetaldehyde acylating dehydrogenase. AroG encodes 3-deoxy-d-arabino-heptulosonate synthase. TyrA encodes chorismate mutase/prephenate dehydratase.

We used a two-step heterologous ethanol utilization pathway to convert ethanol into acetyl–coenzyme A (CoA) without consuming adenosine triphosphate (ATP). This pathway entailed the introduction of acetaldehyde dehydrogenase (encoded by ada gene) from Dickeya zeae and alcohol dehydrogenase (encoded by adh2 gene) from Saccharomyces cerevisiae (Fig. 3), respectively (51). Regarding acetic acid, native E. coli is capable of metabolizing it to produce acetyl-CoA through two pathways: the phosphate acetyltransferase-acetate kinase (Pta-AckA) pathway and acetyl-CoA synthesis (Acs) pathway (55).

To establish the l-tyrosine production pathway, we enhanced the strain through two strategies. First, we deleted two genes, (ΔtyrR) and (ΔpheA), to alleviate transcriptional repression on l-tyrosine production and to eliminate phenylalanine (a by-product) formation, respectively. Second, we overexpressed the feedback-resistant variants of aroG and tyrA for enhanced l-tyrosine production (Fig. 3). As a result, the newly constructed EUTPP strain (strain with ethanol-utilizing and tyrosine-producing pathways) in this study yields l-tyrosine via the metabolism of both acetic acid and ethanol at the same time (Fig. 4, A and B, and fig. S18). In comparison with strain MG1655DE3 [the parent E. coli strain (56)] and strain TpyrA_AH [with only ethanol-utilizing pathway (51)], both strain TPP15 (strain with tyrosine-producing pathway (56)] and strain EUTPP demonstrated effective tyrosine production (Fig. 4B), thanks to their enhanced tyrosine production capabilities. This observation strongly indicates that the presence of the l-tyrosine pathway has a substantial impact on the production of tyrosine (Fig. 3). As anticipated, we observe that both strains of PgyrA_AH and EUTPP exhibited higher rate of ethanol consumption compared to strains of MG1655DE3 and TPP15, thanks to the incorporation of the ethanol-utilizing pathway (fig. S18A). Besides, both strain TPP15 and strain EUTPP showed relatively diminished assimilation effect of acetic acid, likely indicating that the introduction of l-tyrosine producing plasmid and the deletions of ∆pheA and ∆tyrR might have inhibited the utilization of acetic acid to certain extent (fig. S18B). This inhibition could be attributed to the substantial ATP consumption by the acetate utilization pathway, which is likely provided by the generated reduced form of nicotinamide adenine dinucleotide (NADH) and reduced flavin adenine dinucleotide. The enhanced l-tyrosine production competes with cell growth for building blocks such as NADH and ATP. Consequently, the rate of acetate consumption has been negatively influenced (54). Notably, as depicted in Fig. 4A, we found that the strain EUTPP exhibited superior growth rate compared to the strain TPP15 throughout the entire experiment. Consequently, the production of l-tyrosine in newly constructed strain EUTPP suppressed that of strain TPP15 by over 470%, reaching 0.75 g liter−1 on day 7 (Fig. 4B), despite both strain TPP15 and strain EUTPP displaying similar trends in acetic acid utilization rate (fig. S18B). This substantial enhancement in l-tyrosine production strongly suggests the favorable impact of introducing the ethanol utilization pathway in ethanol-containing substrates.

Fig. 4. Metabolic engineering and optimization of fermentation substrates.

(A) Growth rates of the engineered strain EUTPP and other control strains. A mixed substrate containing acetic acid (~10 g liter−1) and ethanol (1.2 g liter−1) was used here. The exact concentrations are provided in fig. S18. OD600, optical density at 600 nm. (B) l-tyrosine production on engineered EUTPP and other strains. (C) l-tyrosine production on engineered strain EUTPP using acetic acid or ethanol as the sole carbon. (D) Substrate utilization of the engineered strain EUTPP, where the substrate contains both 12C acetic acid and 13C-labeled ethanol. (E) Production of 12C-l-tyrosine and 13C-l-tyrosine by engineered EUTPP, M0 and M1 denote for l-tyrosine contains zero and one 13C atom, respectively. (F) Atomic ratio of 13C and 12C in the substrate and final l-tyrosine products. (G) Growth of the engineered strain EUTPP at various substrate concentrations. The acetic acid and ethanol concentration increased from ~2.5 to 20 g liter−1 for substrates 1 to 4, and the exact concentrations are provided in fig. S20. (H) l-tyrosine production on engineered strain EUTPP using various substrate concentrations. (I) l-tyrosine production rate of engineered strain EUTPP using various substrate concentrations.

To investigate the contributions of both acetic acid and ethanol as carbon sources to l-tyrosine production, we first conducted fermentations using the engineered EUTPP strain with acetic acid and ethanol individually as the sole carbon sources. When feeding ethanol as the exclusive carbon source at ~10 g liter−1, the strain EUTPP yielded approximately 1.17 g liter−1 of l-tyrosine by day 7 (fig. S19). Notably, during this process, the strain EUTPP exhibited an obviously accelerated ethanol consumption rate of 5 g liter−1 day−1, outpacing the strain TpyrA_AH’s rate of 2 g liter−1 day−1, which lack the additional l-tyrosine producing pathway (51). By harboring the indigenous capability of E. coli for acetic acid assimilation, strain EUTPP consistently consumed acetic acid (~10 g liter−1, exclusive carbon source; fig. S19) over a span of 7 days at a rate of ~0.8 g liter−1 day−1 and ultimately yielded around 0.58 g liter−1 of l-tyrosine. Nevertheless, even though the rates of carbon source utilization and l-tyrosine production substantially diverged between ethanol and acetic acid as carbon sources, the l-tyrosine production to consumed acetic acid/ethanol mole ratios remained similar (Fig. 4C), indicating that both acetic acid and ethanol are promising feedstocks for l-tyrosine production.

Given that both acetic acid and ethanol can be used as carbon sources in the engineered EUTPP for l-tyrosine production, there is a need for further exploration to comprehend the mechanisms underlying the improvement in l-tyrosine production when introducing the ethanol utilization pathway in COR-derived substrates. To investigate the specific contributions of acetic acid and ethanol to the l-tyrosine production in the engineered EUTPP, we prepared a blended substrate, which consists of 12C acetic acid and 13C2-labeled ethanol, with concentration ratios similar to those in our COR-derived substrate. Under the same fermentation conditions, both 12C acetic acid and 13C ethanol were subsequently consumed, as shown in Fig. 4D. However, an unexpected small amount of 13C was incorporated into the l-tyrosine production. As shown in Fig. 4E, 12C-l-tyrosine (M0) was produced as the predominate product despite that the mass ratio of 13C-labeled ethanol exceeded 10% in the substrates. Furthermore, the atomic ratio of 13C to 12C in the final l-tyrosine products (~2%) was notably lower than that in the initial substrate (Fig. 4F and note S1). Together, we attribute the above performance enhancement to the cooperative interaction between the ethanol pathway and the acetic acid pathway within the EUTPP strain. Because the acetate pathway requires large quantity of ATP, which is provided by oxidizing NADH, oxygen transport would serve as a limiting factor, causing redox imbalance (accumulation of NADH and lack of ATP and NAD+). The ethanol pathway produces an excessive amount of NADH, resulting in less flux into the oxidative TCA cycle, more generated ATP to promote strain metabolism, and thus inducing a higher tyrosine yield. Specifically, we believe that the ethanol pathway primarily contributes to the growth of the strain. In addition to the isotopic labelling experiment, this is also supported by the enhanced growth rate of strain PgyrA_AH compared to other strains (Fig. 4A). Consequently, the enhanced growth of the strain further promotes the acetic acid pathway within the EUTPP strain for the production of l-tyrosine.

We have also explored the optimal concentration of acetic acid and ethanol as co-carbon sources for l-tyrosine production (Fig. 4, G and H, and figs. S20 and S21), aiming to mitigate the inhibitory effects of high substrate concentrations on the growth of E. coli (57–59). Note that the mass ratio of acetic acid to ethanol in the feeds was determined according to the results from our electrochemical COR experiments. At low substrate concentrations, strain EUTPP exhibited steady growth over the course of seven monitoring days. However, the stationary phase was quickly reached on day 1 for a substrate concentration of 2.5 g liter−1 (substrate 1) and on day 2 for a substrate concentration of 5 g liter−1 (substrate 2). The optimal l-tyrosine production was observed at approximately 0.75 g liter−1 when the substrate concentration of 10 g liter−1 (substrate 3) was used. Further increasing the substrate concentration to 20 g liter−1 (substrate 4) resulted in the rapid inhibition and cessation of strain EUTPP growth, as shown in Fig. 4G. This effect was caused by the high concentration acetic acid (fig. S22).

Nevertheless, when using substrate 3, the engineered strain EUTPP achieved an encouraging l-tyrosine production rate of approximately 0.11 g liter−1 day−1 (Fig. 4I). Moreover, we verified that the blended carbon substrates exhibited higher mole ratio of l-tyrosine to consumed substrate (4.07%; fig. S23), compared to the individual carbon substrates (3.25% for ethanol and 3.45% for acetate; Fig. 4C), further confirming the superiority of using the blended carbon substrates for fermentations. In addition, higher ethanol ratio in the blended nexus molecules that exhibit higher l-tyrosine production also confirm the promotion effects of ethanol utilization pathway (fig. S24).

Integrated system optimization

We tackled the challenges presented by the blended electrochemical products and impurities (e.g., high concentration salts) by using genetically engineered strains and electrocatalytic reactors based on solid electrolyte. Nevertheless, the aforementioned electrocatalytic reactor is restricted to producing low-concentration products, thereby limiting l-tyrosine production in the biotic system. As shown in Fig. 4H, the concentration of the fermentation substrate (acetic acid and ethanol) directly affects the production rate of l-tyrosine. In the initial electrocatalytic system design, the electrochemically generated oxygenates initially accumulate at the side of the anion exchange membrane. Subsequently, driven by the concentration gradient and electronic field, they migrate into the solid electrolyte layer. Although the product concentrations can be potentially increased by reducing the water flow rate within the solid electrolyte layer, notable reduction in COR selectivity will occur (46, 60). This is because that elevated local product concentrations can induce changes in the local environment of COR, such as shifts in pH from alkaline to acidic due to the accumulation of the acetic acid. These changes can lead to decreased selectivity (46, 48). To circumvent this challenge, we introduced an approach of recycling water flow within the solid electrolyte layer. By maintaining the water flow rate at 1 ml min−1 and recycling the generated products, we achieved a gradual increase in product concentration. The concentration of acetic acid and ethanol reached 16.5 and 1.3 g liter−1, respectively, after 10-hour operation (fig. S25). The total acetic acid and ethanol FEs of our design (~30%) was higher than the previous single-pass design (~20%) (60). Owing to the relatively high water-flow rate, the generated oxygenated products rapidly diffused into the bulk solution, preventing excessive localized local product concentrations. Consequently, this approach allowed us to produce sufficiently concentrated products while maintaining the high selectivity and stability (46).

Although increased product concentrations can be achieved upon introducing recycling water flow, we observed substantial reduction in the overall FE toward acetic acid and ethanol (i.e., ~30%). We hypothesized that a portion of the generated products may be penetrating through the gas diffusion electrode and subsequently carried away by the gas flow. To validate this, we introduced an extra cold trap along the gas flow path aiming to capture the potentially lost products (Fig. 5A). Notably, this facile addition led to a substantial increase in C2 oxygenates FE to over 40% (Fig. 5B). Through these optimizations, we achieved a product solution containing acetic acid (23.8 g liter−1), ethanol (2.8 g liter−1), and trace amounts of n-propanol (Fig. 5B and fig. S26). This was achieved over a 10-hour electrolysis at 600 mA cm−2 (fig. S26A). As the electrolysis time increased, the concentrations of acetic acid and ethanol rose (fig. S26B), with the solution remaining free from electrolytes, metal impurities (table S2), and active oxygen species (fig. S27). This enabled the direct use of the solution for microbial fermentation without the need for downstream separation or purification processes (table S3). The trace amount of n-propanol exhibited negligible inhibitory effects on l-tyrosine production (fig. S28). While ethanol and acetic acid exhibit comparable mole-normalized l-tyrosine production capabilities (Fig. 4C) and theoretical yield (note S2), it is worth noting that in the electrocatalytic process, the production of one ethanol molecule from CO2 requires 12 electrons, whereas acetic acid only requires 8 electrons. As a result, the acetic acid pathway demonstrates much higher overall electron efficiency, or, in other words, energy efficiency, for l-tyrosine production from CO2 compared to ethanol pathway (Fig. 5C). In light of this, when maintaining a constant current, our preference is for a higher product selectivity of acetic acid. However, a certain amount of ethanol is beneficial for initiating the above positive cooperative effects. Consequently, we have selected an electrolysis current density of 600 mA cm−2. This choice is grounded in its demonstrated high production rate of blended nexus molecules over extended operation times (10 hours) and an optimized product ratio between ethanol and acetate.

Fig. 5. Integrated system optimization for the abiotic/biotic cascade catalysis using blended nexus molecules.

(A) System optimization for high concentration C2 oxygenates production. We recycled the solution in the solid electrolyte layer and added a cold trap to condensate the liquid vapor in the gas phase. (B) Liquid products FEs and liquid products concentration of as-generated products using optimized electrocatalytic system. (C) Comparison of electron efficiency using acetic acid and ethanol (produced from CO2) as feedstock. (D) Acetic acid and ethanol concentrations obtained from COR using different electrolytes. (E) Growth curve of EUTPP strain using different substrates obtained from COR using different electrolytes. (F) Mass spectra recorded for precursor ions at m/z 182.1, corresponding to l-tyrosine produced from the cascade system. (G) l-tyrosine production rate by engineered EUTPP using different substrates obtained from COR using different electrolytes. (H) LC-MS signals of the l-tyrosine produced using different substrates after 7 days. (I) Techno-economic analysis of l-tyrosine production via the cascade abiotic/biotic catalytic system.

As aforementioned, the optimized substrate concentration is approximately 10 g liter−1, for the growth of E. coli. and the productivity of l-tyrosine (Fig. 4, G and H). Provided the potential for producing high concentration acetic acid and ethanol using the modified solid electrolyte reactors, it is straightforward to obtain the desired substrate concentration for microbial fermentation simply via dilution (Fig. 5D). As shown in Fig. 5E, the growth of strain EUTPP using the electrochemically synthesized substrate is evident through the rise in optical density at 600 nm, which increased from 0.05 to 0.97 within a span of 7 days. Besides, the successful production of l-tyrosine using the spatially decoupled biotic/abiotic cascade catalytic system (fig. S29) is confirmed by mass spectra, which exhibits a prominent peak at 182.1 mass/charge ratio (m/z) (Fig. 5F), corresponding to an approximate concentration of 0.6 g liter−1 of l-tyrosine (Fig. 5G). The overall energy efficiency of the abiotic/biotic catalytic system was about 0.41% (note S3). In contrast, when the electrochemically produced acetate and ethanol were mixed with commonly used electrolytes, such as 1.0 M KHCO3 or 1.0 M KOH, only trace amounts of l-tyrosine could be produced by strain EUTPP within the same period (Fig. 5, D to H, and fig. S30). This is likely due to the inhibition of E. coli growth caused by the concentrated electrolyte salts (Fig. 5E).

Using these promising preliminary data, we conducted a preliminary techno-economic analysis (see details in note S4 and tables S4 and S5) to assess the economic feasibility of our cascade abiotic/biotic catalytic process. As shown in Fig. 5I, with ambitious, yet achievable assumptions (electricity price of 0.02 US$ kWh−1), our system demonstrates the potential for l-tyrosine production at a cost (~11,375 US$ ton−1) lower than the current market price (14,740 US$ ton−1) (61). Moreover, the primary factor contributing to the l-tyrosine production cost is the expenditure on electricity. This highlights the need for ongoing research to further enhance oxygenates selectivity and reduce the overpotential for electrochemical CO2/CO conversion, as well as the importance of advancing bioengineered strains to attain higher conversion efficiency.

DISCUSSION

Our study demonstrated a sustainable pathway for synthesizing l-tyrosine from CO2, using a cascade abiotic/biotic catalytic system that uses a blend of nexus molecules. The primary challenge for cascade l-tyrosine production stems from the disparity between the supply of electrocatalysis and the demand for biotic catalysis of nexus molecules, which was addressed through genetic and electrocatalytic engineering. Specifically, electrocatalytic engineering, which incorporates a solid electrolyte reactor and a recycled products flow technique, is used to prevent salt mixture and increase product concentration. A bioengineered E. coli strain, named EUTPP, specifically designed to harbor the ethanol and acetic acid utilization pathway and the l-tyrosine producing pathway. We show that the electrochemically produced mixture of acetic acid and ethanol can be directly used as blended nexus molecules for biofermentation based on EUTPP. Notably, our findings indicate that the incorporation of the ethanol pathway synergies with the preexisting acetic acid–based pathway, leading to a substantially enhanced production rate of l-tyrosine and increased carbon utilization efficiency. Specifically, we have successfully achieved l-tyrosine synthesis at a concentration of 0.6 g liter−1 via this cascade abiotic/biotic catalysis. Furthermore, a preliminary techno-economic assessment has substantiated the economic viability of our cascade abiotic/biotic catalytic system for the production of l-tyrosine. Future research endeavors focusing on both enhancing the performance of electrochemical CO2/CO conversion, as well as improving l-tyrosine production by manipulating the central metabolism and introducing new pathways for acetic acid and ethanol assimilation, can further advancing the sustainable production of l-tyrosine.

MATERIALS AND METHODS

Catalyst synthesis and electrode preparation

We used a same method to synthesis Cu, Ag, and Cu-Ag nanoparticles. First, x (0 ≤ x ≤ 10) mmol Cu(NO3)2 and 10 − x mmol AgNO3 were dissolved in 10 ml of DI water, and the mixture was heated to 90°C. Then, 3 g of KOH and 1.5 g of K2S2O8 were dissolved in 50 ml of DI water and drop added to above solution. After 1 hour, the solid powder was obtained by centrifugation and washed using ethanol and water for three times and dried in vacuum. Last, before COR test, the catalyst was in situ reduced to metallic species at 200 mA cm−2. To prepare the cathode ink, 40-mg catalyst with 160 μl of Nafion D520 polymer binder solution was suspended in 8 ml of isopropyl alcohol. Around 0.7 mg cm−2 of catalyst was loaded on gas diffusion layer electrode as the cathode electrode.

Characterization

Scanning electron microscopy (SEM) images were captured by a JEOL, JSM-7610F SEM. TEM images were captured by a JEOL JEM-2010 TEM (working voltage, 200 kV). The XRD spectra was carried out by a GBC MMA x-ray diffractometer with a Cu Kα irradiation source (λ = 1.54056 Å). The XPS spectra were tested using a Thermo ESCALAB 250 (monochrome Al Kα, hν = 1486.6 eV), and the calibration was done by setting the C 1s peak at 284.6 eV. Perkin Elmer Avio 500 inductively coupled plasma-optical emission spectrometer was used for element concentration test. Fluorescence spectra were recorded using a FluoroMax-4 spectrofluorometer.

Electrochemical measurement

A BioLogic VMP3 workstation was used to record the electrochemical response. The overall resistance was determined by potentiostatic electrochemical impedance spectroscopy at frequencies ranging from 0.1 Hz to 200 kHz. All the measured potentials and voltages were manually compensated. The typical three-electrode measurements were performed at 25°C using a customized flow cell separated by Nafion 117 membrane. The geometry surface area of the cathode electrode was fixed to 1 cm2. The anode side was circulated with 1 M KOH aqueous solution at 1 ml min−1. The anode side was circulated with 1 M H2SO4 aqueous solution at 2 ml min−1. A IrO2-coated Ti mesh and a saturated calomel electrode (SCE; CH Instruments) were used as the counter and reference electrodes, respectively. All potentials measured against SCE (ESCE) were converted to the RHE (ERHE) scale in this work using ERHE = ESCE + 0.244 V + 0.0591 × pH, where pH values of the electrolytes were determined by a pH meter.

For the two-electrode cell using solid electrolytes, an anion exchange membrane (Dioxide Materials) and a Nafion film (Fuel Cell Store) were used for anion and cation exchange, respectively. An IrO2-coated Ti mesh was used as the anode electrodes. The cathode side was supplied with 20 standard cubic centimeters per minute (sccm; monitored by Alicat Scientific mass flow controller) CO gas. The water flow rate in the solid-state electrolyte layer was 1 ml min−1. The anode side was circulated with 1 M H2SO4 aqueous solution at 2 ml min−1. The porous styrene-divinylbenzene sulfonated copolymer was used as solid ion conductors. To produce feedstock for the fermentation, the solid electrolyte reactor was operated under a current density of 600 mA cm−2, and 45 ml of DI water was cycled through the solid electrolyte for 10 hours. For the membrane electrode assembly test for CO2RR, an anion exchange membrane (Dioxide Materials) was used for anion exchange. An IrO2-coated Ti mesh was used as the anode electrodes. The cathode side was supplied with 20-sccm CO2 gas. The anode side was circulated with 1 M KOH aqueous solution at 2 ml min−1. All the measured potentials using two-electrode setup were manually compensated. Gas chromatograph (Shimadzu GC-2014) with thermal conductivity detector and flame ionization detector with a methanizer was used to quantitative analysis the CO2RR products. One-dimensional 1H NMR spectra were collected on a Bruker 400-MHz spectrometer to quantify the liquid products. Typically, 600 μl of electrolyte after electrolysis were mixed with 100 μl of D2O and 0.05 μl of dimethyl sulfoxide as internal standard.

Strain construction

Plasmids carrying ethanol-utilizing pathway (51) and l-tyrosine–producing pathway (56) were constructed according to guanin/thymine standard (62) in previous studies (table S6). Both of the plasmids were simultaneously introduced into E. coli MG1655DE3 to derive a strain coupling the utilization of ethanol with l-tyrosine production using the standard heat-shock method. The successfully constructed strain were then isolated on LB agar plate containing antibiotics, both ampicillin (25 mg liter−1) and spectinomycin (50 mg liter−1), as selective pressure, dubbed as EUTPP.

Growth medium and culture conditions

To investigate the kinetics of constructed strain EUTPP, strain EUTPP was inoculated (2%, v/v) into 100-ml flasks containing 10 ml of LB medium spiked with spectinomycin (50 mg liter−1) and ampicillin (25 mg liter−1). In addition to antibiotics, the LB medium was also amended with Complete Supplement Mixture (1 g liter−1; Sunrise Science, Singapore), M9 medium (11.3 g liter−1; Bio Basic Asia Pacific Pte Ltd., Singapore), and a mineral salt K3 solution, NH4Cl (2 g liter−1) as previously described (51, 56, 63), as well as isopropyl β-d-1-thiogalactopyranoside (0.1 mM) to induce the expression of promoter (P_lac). Acetic acid and ethanol were added as the carbon source either solely or simultaneously. All the experimental were incubated at 30°C and 250 rpm. Samplings were conducted at interval of 24 hours and stored in −80°C for further analysis. During cultivation, pH was monitored daily and manually adjusted by hydrochloride acid at 7.0 to 8.0. All the chemicals were purchased from Sigma-Aldrich unless specifically mentioned.

Cell growth and analysis of substrates and metabolites

Cell growth was monitored using a Biospec-1601 spectrophotometer (Shimadzu Co., Kyoto, Japan) at 600 nm. The collected samples were centrifuged at 12000 rpm for 10 min and filtered 0.22-μm syringe filter for further chemical analysis. Acetic and ethanol consumption were monitored by Agilent Gas Chromatograph 7890A equipped with a flame ionization detector and a DB-WAXetr column (J&W Scientific, Folsom, CA, USA) as previously described (51). l-tyrosine was 100× diluted by 0.1% formic acid and measured using a liquid chromatography–mass spectrometry (LC-MS) with a column of Agilent ZORBAX 478 Eclipse plus C18 (3.5 μm, 4.6 × 100 mm) with a co-elute of 10% (v/v) acetonitrile and 0.1% (v/v) trifluoracetic acid at 30°C. All the data were biological replicates for two times.

Acknowledgments

We acknowledge V. F. K. Yuen for the help of LC-MS test.

Funding: We acknowledge the National University of Singapore, Ministry of Education for their financial support, through the grants of A-0009176-02-00 and A-0009176-03-00. L.W. acknowledges the support by National Research Foundation (NRF) Singapore, under its NRF Fellowship (NRF-NRFF13-2021-0007) and CRP (NRF-CRP27-2021-0004), and A*STAR (Agency for Science, Technology and Research) under its LCERFI program award no. U2102d2002, as well as the support from the Centre for Hydrogen Innovations at the NUS (CHI-P2022-06).

Author contributions: Conceptualization: L.F., Z.Z., S.P., Jingyi C., L.C., J.H., K.Z., and L.W. Methodology: L.F., Z.Z., S.Z., S.P., Y.Z., L.C., Junmei C., J.H., and L.W. Investigation: L.F., Z.Z., S.P., L.C., and L.W. Resources: L.F., Z.Z., S.P., J.H., and L.W. Funding acquisition: J.H. and L.W. Data curation: Z.Z., J.H., and L.W. Validation: L.F., Z.Z., S.P., Y.Z., L.C., Junmei C., J.H., and L.W. Formal analysis: L.F., Z.Z., S.Z., Y.Z., L.C., and L.W. Project administration: S.Z., S.P., J.H., and L.W. Visualization: L.F., Z.Z., S.P., Y.Z., Jingyi C., L.C., J.H., and L.W. Writing–original draft: L.F., Z.Z., Y.Z., Jingyi C., and L.W. Writing–review and editing: L.F., Z.Z., S.P., Y.Z., Jingyi C., J.H., K.Z, and L.W. Supervision: J.H., K.Z, and L.W.

Competing Interests: The authors declare that they have no competing interests.

Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials.

Supplementary Materials

This PDF file includes:

Figs. S1 to S30

Notes S1 to S4

Tables S1 to S6

References
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