Evolution-assisted engineering of E. coli enables growth on formic acid at ambient CO2 via the Serine Threonine Cycle.

Atmospheric CO2 poses a major threat to life on Earth by causing global warming and climate change. On the other hand, it can be considered as a resource that is scalable enough to establish a circular carbon economy. Accordingly, technologies to capture and convert CO2 into reduced one-carbon (C1) compounds (e.g. formic acid) are developing and improving fast. Driven by the idea of creating sustainable bioproduction platforms, natural and synthetic C1-utilization pathways are engineered into industrially relevant microbes. The realization of synthetic C1-assimilation cycles in living organisms is a promising but challenging endeavour. Here, we engineer the Serine Threonine Cycle, a synthetic C1-assimilation cycle in Escherichia coli to achieve growth on formic acid. Our stepwise engineering approach in tailored selection strains combined with adaptive laboratory evolution experiments enabled formatotrophic growth of the organism. Whole genome sequencing and reverse engineering allowed us to determine the key mutations linked to pathway activity. The Serine Threonine Cycle strains created in this work use formic acid as a carbon and energy source and can grow at ambient CO2 cultivation conditions. This work sets an example for the engineering of complex C1-assimilation cycles in heterotrophic microbes.

Strangers in a foreign land: 'Yeastizing' plant enzymes.

Expressing plant metabolic pathways in microbial platforms is an efficient, cost-effective solution for producing many desired plant compounds. As eukaryotic organisms, yeasts are often the preferred platform. However, expression of plant enzymes in a yeast frequently leads to failure because the enzymes are poorly adapted to the foreign yeast cellular environment. Here, we first summarize the current engineering approaches for optimizing performance of plant enzymes in yeast. A critical limitation of these approaches is that they are labour-intensive and must be customized for each individual enzyme, which significantly hinders the establishment of plant pathways in cellular factories. In response to this challenge, we propose the development of a cost-effective computational pipeline to redesign plant enzymes for better adaptation to the yeast cellular milieu. This proposition is underpinned by compelling evidence that plant and yeast enzymes exhibit distinct sequence features that are generalizable across enzyme families. Consequently, we introduce a data-driven machine learning framework designed to extract 'yeastizing' rules from natural protein sequence variations, which can be broadly applied to all enzymes. Additionally, we discuss the potential to integrate the machine learning model into a full design-build-test cycle.

Engineering new-to-nature biochemical conversions by combining fermentative metabolism with respiratory modules.

Anaerobic microbial fermentations provide high product yields and are a cornerstone of industrial bio-based processes. However, the need for redox balancing limits the array of fermentable substrate-product combinations. To overcome this limitation, here we design an aerobic fermentative metabolism that allows the introduction of selected respiratory modules. These can use oxygen to re-balance otherwise unbalanced fermentations, hence achieving controlled respiro-fermentative growth. Following this design, we engineer and characterize an obligate fermentative Escherichia coli strain that aerobically ferments glucose to stoichiometric amounts of lactate. We then re-integrate the quinone-dependent glycerol 3-phosphate dehydrogenase and demonstrate glycerol fermentation to lactate while selectively transferring the surplus of electrons to the respiratory chain. To showcase the potential of this fermentation mode, we direct fermentative flux from glycerol towards isobutanol production. In summary, our design permits using oxygen to selectively re-balance fermentations. This concept is an advance freeing highly efficient microbial fermentation from the limitations imposed by traditional redox balancing.

Automated in vivo enzyme engineering accelerates biocatalyst optimization.

Achieving cost-competitive bio-based processes requires development of stable and selective biocatalysts. Their realization through in vitro enzyme characterization and engineering is mostly low throughput and labor-intensive. Therefore, strategies for increasing throughput while diminishing manual labor are gaining momentum, such as in vivo screening and evolution campaigns. Computational tools like machine learning further support enzyme engineering efforts by widening the explorable design space. Here, we propose an integrated solution to enzyme engineering challenges whereby ML-guided, automated workflows (including library generation, implementation of hypermutation systems, adapted laboratory evolution, and in vivo growth-coupled selection) could be realized to accelerate pipelines towards superior biocatalysts.

Prospects of formamide as nitrogen source in biotechnological production processes.

This review presents an analysis of formamide, focussing on its occurrence in nature, its functional roles, and its promising applications in the context of the bioeconomy. We discuss the utilization of formamide as an innovative nitrogen source achieved through metabolic engineering. These approaches underscore formamide's potential in supporting growth and production in biotechnological processes. Furthermore, our review illuminates formamide's role as a nitrogen source capable of safeguarding cultivation systems against contamination in non-sterile conditions. This attribute adds an extra layer of practicality to its application, rendering it an attractive candidate for sustainable and resilient industrial practices. Additionally, the article unveils the versatility of formamide as a potential carbon source that could be combined with formate or CO2 assimilation pathways. However, its attributes, i.e., enriched nitrogen content and comparatively limited energy content, led to conclude that formamide is more suitable as a co-substrate and that its use as a sole source of carbon for biomass and bio-production is limited. Through our exploration of formamide's properties and its applications, this review underscores the significance of formamide as valuable resource for a large spectrum of industrial applications. KEY POINTS: • Formidases enable access to formamide as source of nitrogen, carbon, and energy • The formamide/formamidase system supports non-sterile fermentation • The nitrogen source formamide supports production of nitrogenous compounds.

Engineering a synthetic energy-efficient formaldehyde assimilation cycle in Escherichia coli.

One-carbon (C1) substrates, such as methanol or formate, are attractive feedstocks for circular bioeconomy. These substrates are typically converted into formaldehyde, serving as the entry point into metabolism. Here, we design an erythrulose monophosphate (EuMP) cycle for formaldehyde assimilation, leveraging a promiscuous dihydroxyacetone phosphate dependent aldolase as key enzyme. In silico modeling reveals that the cycle is highly energy-efficient, holding the potential for high bioproduct yields. Dissecting the EuMP into four modules, we use a stepwise strategy to demonstrate in vivo feasibility of the modules in E. coli sensor strains with sarcosine as formaldehyde source. From adaptive laboratory evolution for module integration, we identify key mutations enabling the accommodation of the EuMP reactions with endogenous metabolism. Overall, our study demonstrates the proof-of-concept for a highly efficient, new-to-nature formaldehyde assimilation pathway, opening a way for the development of a methylotrophic platform for a C1-fueled bioeconomy in the future.

The cofactor challenge in synthetic methylotrophy: bioengineering and industrial applications.

Methanol is a promising feedstock for industrial bioproduction: it can be produced renewably and has high solubility and limited microbial toxicity. One of the key challenges for its bio-industrial application is the first enzymatic oxidation step to formaldehyde. This reaction is catalysed by methanol dehydrogenases (MDH) that can use NAD+, O2 or pyrroloquinoline quinone (PQQ) as an electron acceptor. While NAD-dependent MDH are simple to express and have the highest energetic efficiency, they exhibit mediocre kinetics and poor thermodynamics at ambient temperatures. O2-dependent methanol oxidases require high oxygen concentrations, do not conserve energy and thus produce excessive heat as well as toxic H2O2. PQQ-dependent MDH provide a good compromise between energy efficiency and good kinetics that support fast growth rates without any drawbacks for process engineering. Therefore, we argue that this enzyme class represents a promising solution for industry and outline engineering strategies for the implementation of these complex systems in heterologous hosts.

Formamide-based production of amines by metabolically engineering Corynebacterium glutamicum.

Formamide is rarely used as nitrogen source by microorganisms. Therefore, formamide and formamidase have been used as protection system to allow for growth under non-sterile conditions and for non-sterile production of acetoin, a product lacking nitrogen. Here, we equipped Corynebacterium glutamicum, a renowned workhorse for industrial amino acid production for 60 years, with formamidase from Helicobacter pylori 26695, enabling growth with formamide as sole nitrogen source. Thereupon, the formamide/formamidase system was exploited for efficient formamide-based production of the nitrogenous compounds L-glutamate, L-lysine, N-methylphenylalanine, and dipicolinic acid by transfer of the formamide/formamidase system to established producer strains. Stable isotope labeling verified the incorporation of nitrogen from formamide into biomass and the representative product L-lysine. Moreover, we showed ammonium leakage during formamidase-based access of formamide to be exploitable to support growth of formamidase-deficient C. glutamicum in co-cultivation and demonstrated that efficient utilization of formamide as sole nitrogen source benefitted from overexpression of formate dehydrogenase. KEY POINTS: • C. glutamicum was engineered to access formamide. • Formamide-based production of nitrogenous compounds was established. • Nitrogen cross-feeding supported growth of a formamidase-negative strain.

Implementation of the ß-hydroxyaspartate cycle increases growth performance of Pseudomonas putida on the PET monomer ethylene glycol.

Ethylene glycol (EG) is a promising next generation feedstock for bioprocesses. It is a key component of the ubiquitous plastic polyethylene terephthalate (PET) and other polyester fibers and plastics, used in antifreeze formulations, and can also be generated by electrochemical conversion of syngas, which makes EG a key compound in a circular bioeconomy. The majority of biotechnologically relevant bacteria assimilate EG via the glycerate pathway, a wasteful metabolic route that releases CO2 and requires reducing equivalents as well as ATP. In contrast, the recently characterized ß-hydroxyaspartate cycle (BHAC) provides a more efficient, carbon-conserving route for C2 assimilation. Here we aimed at overcoming the natural limitations of EG metabolism in the industrially relevant strain Pseudomonas putida KT2440 by replacing the native glycerate pathway with the BHAC. We first prototyped the core reaction sequence of the BHAC in Escherichia coli before establishing the complete four-enzyme BHAC in Pseudomonas putida. Directed evolution on EG resulted in an improved strain that exhibits 35% faster growth and 20% increased biomass yield compared to a recently reported P. putida strain that was evolved to grow on EG via the glycerate pathway. Genome sequencing and proteomics highlight plastic adaptations of the genetic and metabolic networks in response to the introduction of the BHAC into P. putida and identify key mutations for its further integration during evolution. Taken together, our study shows that the BHAC can be utilized as 'plug-and-play' module for the metabolic engineering of two important microbial platform organisms, paving the way for multiple applications for a more efficient and carbon-conserving upcycling of EG in the future.

Optimizing E. coli as a formatotrophic platform for bioproduction via the reductive glycine pathway.

Microbial C1 fixation has a vast potential to support a sustainable circular economy. Hence, several biotechnologically important microorganisms have been recently engineered for fixing C1 substrates. However, reports about C1-based bioproduction with these organisms are scarce. Here, we describe the optimization of a previously engineered formatotrophic Escherichia coli strain. Short-term adaptive laboratory evolution enhanced biomass yield and accelerated growth of formatotrophic E. coli to 3.3 g-CDW/mol-formate and 6 h doubling time, respectively. Genome sequence analysis revealed that manipulation of acetate metabolism is the reason for better growth performance, verified by subsequent reverse engineering of the parental E. coli strain. Moreover, the improved strain is capable of growing to an OD600 of 22 in bioreactor fed-batch experiments, highlighting its potential use for industrial bioprocesses. Finally, demonstrating the strain's potential to support a sustainable, formate-based bioeconomy, lactate production from formate was engineered. The optimized strain generated 1.2 mM lactate -10% of the theoretical maximum- providing the first proof-of-concept application of the reductive glycine pathway for bioproduction.

Integrated rational and evolutionary engineering of genome-reduced Pseudomonas putida strains promotes synthetic formate assimilation.

Formate is a promising, water-soluble C1 feedstock for biotechnology that can be efficiently produced from CO2-but formatotrophy has been engineered in only a few industrially-relevant microbial hosts. We addressed the challenge of expanding the feedstock range of bacterial hosts by adopting Pseudomonas putida as a robust platform for synthetic formate assimilation. Here, the metabolism of a genome-reduced variant of P. putida was radically rewired to establish synthetic auxotrophies that could be functionally complemented by expressing components of the reductive glycine (rGly) pathway. We adopted a modular engineering approach, dividing C1 assimilation in segments composed of both heterologous activities (sourced from Methylobacterium extorquens) and native biochemical reactions. Modular expression of rGly pathway elements enabled growth on formate as carbon source and acetate (predominantly for energy supply), and adaptive laboratory evolution of two lineages of engineered P. putida formatotrophs lead to doubling times of ca. 15 h. We likewise identified emergent metabolic features for assimilation of C1 units in these evolved P. putida populations. Taken together, our results consolidate the landscape of useful microbial platforms that can be implemented for C1-based biotechnological production towards a formate bioeconomy.

Optimizing microbial networks through metabolic bypasses.

Metabolism has long been considered as a relatively stiff set of biochemical reactions. This somewhat outdated and dogmatic view has been challenged over the last years, as multiple studies exposed unprecedented plasticity of metabolism by exploring rational and evolutionary modifications within the metabolic network of cell factories. Of particular importance is the emergence of metabolic bypasses, which consist of enzymatic reaction(s) that support unnatural connections between metabolic nodes. Such novel topologies can be generated through the introduction of heterologous enzymes or by upregulating native enzymes (sometimes relying on promiscuous activities thereof). Altogether, the adoption of bypasses resulted in an expansion in the capacity of the host's metabolic network, which can be harnessed for bioproduction. In this review, we discuss modifications to the canonical architecture of central carbon metabolism derived from such bypasses towards six optimization purposes: stoichiometric gain, overcoming kinetic limitations, solving thermodynamic barriers, circumventing toxic intermediates, uncoupling product synthesis from biomass formation, and altering redox cofactor specificity. The metabolic costs associated with bypass-implementation are likewise discussed, including tailoring their design towards improving bioproduction.

On the flexibility of the cellular amination network in E coli.

Ammonium (NH4+) is essential to generate the nitrogenous building blocks of life. It gets assimilated via the canonical biosynthetic routes to glutamate and is further distributed throughout metabolism via a network of transaminases. To study the flexibility of this network, we constructed an Escherichia coli glutamate auxotrophic strain. This strain allowed us to systematically study which amino acids serve as amine sources. We found that several amino acids complemented the auxotrophy either by producing glutamate via transamination reactions or by their conversion to glutamate. In this network, we identified aspartate transaminase AspC as a major connector between many amino acids and glutamate. Additionally, we extended the transaminase network by the amino acids ß-alanine, alanine, glycine, and serine as new amine sources and identified d-amino acid dehydrogenase (DadA) as an intracellular amino acid sink removing substrates from transaminase reactions. Finally, ammonium assimilation routes producing aspartate or leucine were introduced. Our study reveals the high flexibility of the cellular amination network, both in terms of transaminase promiscuity and adaptability to new connections and ammonium entry points.

Nitrogen is an essential part of many of the cell's building blocks, including amino acids and nucleotides, which form proteins and DNA respectively. Therefore, nitrogen has to be available to cells so that they can survive and grow. In nature, some microorganisms convert the gaseous form of nitrogen into ammonium, which then acts as the nitrogen source of most organisms, including bacteria, plants and animals. Once cells take up ammonium, it is 'fixed' by becoming part of an amino acid called glutamate, which has a so-called 'amine group' that contains a nitrogen. Glutamate then becomes the central source for passing these amines on to other molecules, distributing nitrogen throughout the cell. This coupling between ammonium fixation and glutamate production evolved over millions of years and occurs in all organisms. However, the complete metabolic network that underlies the distribution of amines remains poorly understood despite decades of research. Furthermore, it is not clear whether ammonium can be fixed in a way that is independent of glutamate. To answer these questions, Schulz-Mirbach et al. used genetic engineering to create a strain of the bacterium E. coli that was unable to make glutamate. These mutant cells could only grow in the presence of certain amino acids, which acted as alternative amine sources. Schulz-Mirbach et al. found that enzymes called transaminases, and one called AspC in particular, were required for the cells to be able to produce glutamate using the amine groups from other amino acids. Notably, Schulz-Mirbach et al. showed that AspC, which had previously been shown to use an amino acid called aspartate as a source of amine groups, is indispensable if the cell is to use the amine groups from other amino acids ­ including histidine, tyrosine, phenylalanine, tryptophan, methionine, isoleucine and leucine. Schulz-Mirbach et al. also discovered that if they engineered the E. coli cells to produce transaminases from other species, the repertoire of molecules that the cells could use as the source of amines to generate glutamate increased. In a final set of experiments, Schulz-Mirbach et al. were able to engineer the cells to fix ammonium by producing aspartate and leucine, thus entirely bypassing the deleted routes of glutamate synthesis. These data suggest that fixing ammonium and distributing nitrogen in E. coli can be very flexible. The results from these experiments may shed light on how cells adapt when there is not a lot of ammonium available. Moreover, this study could advance efforts at metabolic engineering, for example, to create molecules through new pathways or to boost the production of amino acids needed for industrial purposes.

Engineering the Reductive Glycine Pathway: A Promising Synthetic Metabolism Approach for C1-Assimilation.

In recent years the reductive glycine pathway (rGlyP) has emerged as a promising pathway for the assimilation of formate and other sustainable C1-feedstocks for future biotechnology. It was originally proposed as an attractive "synthetic pathway" to support formatotrophic growth due to its high ATP efficiency, linear structure, and limited overlap with native pathways in most microbial hosts. Here, we present the current state of research on this pathway including breakthroughs on its engineering. Different variants of the rGlyP are discussed, including its core module for formate to glycine conversion, as well as varying modules for substrate conversion to formate, and glycine assimilation routes. Very recently, the rGlyP has been successfully implemented for synthetic formatotrophic growth, as well as for growth on methanol, in some bacterial hosts. We discuss the engineering strategies employed in these studies, including growth-coupled selection of functional pathway modules. We also compare the rGlyP to other natural and synthetic C1-assimilation pathways. Finally, we provide an outlook on open challenges and opportunities for the rGlyP, including its engineering into more biotechnological hosts, as well as the still-to-be realized production of value-added chemicals via this pathway. We expect that further research on the rGlyP will support the efficient use of sustainable C1-substrates in bioproduction.

Toward a glycyl radical enzyme containing synthetic bacterial microcompartment to produce pyruvate from formate and acetate.

Formate has great potential to function as a feedstock for biorefineries because it can be sustainably produced by a variety of processes that don't compete with agricultural production. However, naturally formatotrophic organisms are unsuitable for large-scale cultivation, difficult to engineer, or have inefficient native formate assimilation pathways. Thus, metabolic engineering needs to be developed for model industrial organisms to enable efficient formatotrophic growth. Here, we build a prototype synthetic formate utilizing bacterial microcompartment (sFUT) encapsulating the oxygen-sensitive glycyl radical enzyme pyruvate formate lyase and a phosphate acyltransferase to convert formate and acetyl-phosphate into the central biosynthetic intermediate pyruvate. This metabolic module offers a defined environment with a private cofactor coenzyme A that can cycle efficiently between the encapsulated enzymes. To facilitate initial design-build-test-refine cycles to construct an active metabolic core, we used a "wiffleball" architecture, defined as an icosahedral bacterial microcompartment (BMC) shell with unoccupied pentameric vertices to freely permit substrate and product exchange. The resulting sFUT prototype wiffleball is an active multi enzyme synthetic BMC functioning as platform technology.

Activating Silent Glycolysis Bypasses in Escherichia coli.

All living organisms share similar reactions within their central metabolism to provide precursors for all essential building blocks and reducing power. To identify whether alternative metabolic routes of glycolysis can operate in E. coli, we complementarily employed in silico design, rational engineering, and adaptive laboratory evolution. First, we used a genome-scale model and identified two potential pathways within the metabolic network of this organism replacing canonical Embden-Meyerhof-Parnas (EMP) glycolysis to convert phosphosugars into organic acids. One of these glycolytic routes proceeds via methylglyoxal and the other via serine biosynthesis and degradation. Then, we implemented both pathways in E. coli strains harboring defective EMP glycolysis. Surprisingly, the pathway via methylglyoxal seemed to immediately operate in a triosephosphate isomerase deletion strain cultivated on glycerol. By contrast, in a phosphoglycerate kinase deletion strain, the overexpression of methylglyoxal synthase was necessary to restore growth of the strain. Furthermore, we engineered the "serine shunt" which converts 3-phosphoglycerate via serine biosynthesis and degradation to pyruvate, bypassing an enolase deletion. Finally, to explore which of these alternatives would emerge by natural selection, we performed an adaptive laboratory evolution study using an enolase deletion strain. Our experiments suggest that the evolved mutants use the serine shunt. Our study reveals the flexible repurposing of metabolic pathways to create new metabolite links and rewire central metabolism.

Change in Cofactor Specificity of Oxidoreductases by Adaptive Evolution of an Escherichia coli NADPH-Auxotrophic Strain.

The nicotinamide cofactor specificity of enzymes plays a key role in regulating metabolic processes and attaining cellular homeostasis. Multiple studies have used enzyme engineering tools or a directed evolution approach to switch the cofactor preference of specific oxidoreductases. However, whole-cell adaptation toward the emergence of novel cofactor regeneration routes has not been previously explored. To address this challenge, we used an Escherichia coli NADPH-auxotrophic strain. We continuously cultivated this strain under selective conditions. After 500 to 1,100 generations of adaptive evolution using different carbon sources, we isolated several strains capable of growing without an external NADPH source. Most isolated strains were found to harbor a mutated NAD+-dependent malic enzyme (MaeA). A single mutation in MaeA was found to switch cofactor specificity while lowering enzyme activity. Most mutated MaeA variants also harbored a second mutation that restored the catalytic efficiency of the enzyme. Remarkably, the best MaeA variants identified this way displayed overall superior kinetics relative to the wild-type variant with NAD+. In other evolved strains, the dihydrolipoamide dehydrogenase (Lpd) was mutated to accept NADP+, thus enabling the pyruvate dehydrogenase and 2-ketoglutarate dehydrogenase complexes to regenerate NADPH. Interestingly, no other central metabolism oxidoreductase seems to evolve toward reducing NADP+, which we attribute to several biochemical constraints, including unfavorable thermodynamics. This study demonstrates the potential and biochemical limits of evolving oxidoreductases within the cellular context toward changing cofactor specificity, further showing that long-term adaptive evolution can optimize enzyme activity beyond what is achievable via rational design or directed evolution using small libraries. IMPORTANCE In the cell, NAD(H) and NADP(H) cofactors have different functions. The former mainly accepts electrons from catabolic reactions and carries them to respiration, while the latter provides reducing power for anabolism. Correspondingly, the ratio of the reduced to the oxidized form differs for NAD+ (low) and NADP+ (high), reflecting their distinct roles. We challenged the flexibility of E. coli's central metabolism in multiple adaptive evolution experiments using an NADPH-auxotrophic strain. We found several mutations in two enzymes, changing the cofactor preference of malic enzyme and dihydrolipoamide dehydrogenase. Upon deletion of their corresponding genes we performed additional evolution experiments which did not lead to the emergence of any additional mutants. We attribute this restricted number of mutational targets to intrinsic thermodynamic barriers; the high ratio of NADPH to NADP+ limits metabolic redox reactions that can regenerate NADPH, mainly by mass action constraints.