Branched-chain fatty acids (BCFAs), naturally synthesized by Gram-positive bacteria, are promising feedstocks for the production of advanced biofuels. However, efficient BCFA biosynthesis in Gram-negative bacteria such as Escherichia coli remains challenging because of insufficient supply of branched-chain acyl-CoA precursors, poor compatibility of the endogenous fatty acid synthesis pathway with branched-chain substrates, and limited cellular robustness toward non-native fatty acids. In this study, we first engineered an orthogonal isovaleryl-CoA biosynthetic pathway in E. coli and demonstrated its functionality in supporting BCFA production. Importantly, we identified a strong synergistic interaction between the isovaleryl-CoA pathway and the branched-chain α-keto acid dehydrogenase pathway, and thereby established a dual-route strategy for precursor supply. To further enhance BCFA production, we rewired central carbon metabolism by eliminating competing pathways and introducing a non-oxidative glycolysis pathway to increase acetyl-CoA availability while minimizing byproduct formation. We additionally optimized the fatty acid biosynthetic module through expression of a highly active 'TesA (R65C) variant and the transcriptional regulator FadR, and enhanced cellular robustness via introduction of N138H mutation into PcnB and overexpression of the stress resistance associated genes rfaY and yafL. The final engineered strain produced 2.96 g/L BCFAs (approximately 10-fold higher than the previously reported titers), representing 55% of total fatty acids, with a yield of 0.08 g/g glucose. Overall, this work established a dual-precursor supply strategy combined with systems metabolic engineering for BCFA production, providing a foundation for the development of sustainable bioprocesses for advanced branched-chain biofuels.

As a high-value-added amino acid derivative, trans-4-hydroxy-L-proline (T-4-Hyp) faces key bottlenecks in its microbial production from glucose, including insufficient precursor supply and an imbalance between cell growth and product biosynthesis. In this study, we successfully constructed an engineered Escherichia coli strain QF-27 for efficient T-4-Hyp production. First, the L-proline (L-Pro) biosynthetic pathway was enhanced by overexpressing the feedback-resistant γ-glutamyl kinase, glutamate-γ-semialdehyde dehydrogenase, and pyrroline-5-carboxylate reductase, and by knocking out the L-proline dehydrogenase. The resulting strain QF-9 produced 15.75 ± 0.56 g/L of L-Pro. Subsequently, the expression level of proline-4-hydroxylase from Dactylosporangium sp. RH1 was optimized in strain QF-9, and the resulting strain QF-14 produced 5.32 ± 0.26 g/L of T-4-Hyp. To address the insufficient supply of α-ketoglutarate (α-KG), a multi-modular synergistic strategy (i.e., blocking byproduct pathways, enhancing α-KG flux, and relieving global transcriptional repression) increased T-4-Hyp production to 11.02 ± 0.27 g/L. Moreover, a dynamic switch combining PrpsT promoter and DAS+4 degradation tag was designed to repress expression and promote degradation of the α-ketoglutarate dehydrogenase complex during the stationary phase, thereby balancing cell growth and T-4-Hyp production. Consequently, T-4-Hyp production reached 14.37 ± 0.46 g/L, and residual L-Pro fell to 0.38 ± 0.17 g/L. In fed-batch fermentation, the final strain QF-27 produced 105.72 ± 0.84 g/L of T-4-Hyp with productivity of 2.20 g/L/h and carbon yield of 0.364 g/g glucose. To our knowledge, this is the best performance reported for T-4-Hyp production by microbial fermentation, and the first study to enhance it via systematic modification of central carbon metabolism.

Tricyclic sesquiterpenes are bioactive natural products with broad applications in pharmaceuticals, fragrances, and sustainable aviation biofuels. However, the repertoire of characterized tricyclic sesquiterpene synthases (tSTSs) remains limited. To systematically expand this enzyme class, we developed a multi-step bioinformatics workflow to identify genes encoding for cyclic sesquiterpene synthase integrating BLAST-based homology filtering, conserved motif validation, AlphaFold2-based structural modeling, and molecular docking. This workflow progressively refined an initial set of 1063 tSTS candidates to six putative enzymes that satisfied a C1-C10 distance criterion. Heterologous expression of all six candidates in Corynebacterium glutamicum JP-2 overexpressing a methylerythritol 4-phosphate pathway gene module confirmed biosynthetic activity, yielding five known cyclic sesquiterpenes and tricyclic avermitilol produced by a tSTS (AtTPS) identified from Actinokineospora terrae by NMR spectroscopy and high-resolution electrospray ionization mass spectrometry. To improve industrial feasibility, the chemical-inducible promoter was replaced with constitutive leaderless synthetic σ[B] promoters, and fusion of avermitilol synthase with farnesyl pyrophosphate synthase via a GGGGS linker enhanced the local concentration of intermediates between enzymes. In addition, a plasmid-free, antibiotic selection-free strain was constructed via the CRISPR-associated transposons, enabling dual chromosomal integrations of the avermitilol synthase expression cassette and achieving 81.52 mg/L avermitilol production. The chromosomally integrated strain maintained stable production over serial passages, whereas the plasmid-based strain exhibited greater than 90% loss of productivity. Fed-batch fermentation of AVM-int02cp in a 2-L bioreactor achieved a final avermitilol titer of 100.99 mg/L. This work demonstrates a sequence- and 3D structure-assisted gene discovery-to-production workflow for a tricyclic sesquiterpene through the identification of an uncharacterized synthase gene in a microbial host.

Human N-glycoproteins represent a market worth hundreds of billions of dollars, yet their production in yeast is often limited by misfolding and degradation. However, few strategies have addressed this limitation by targeting differences between human and yeast N-glycan-dependent protein quality control (QC), including the absence of the UGGT-mediated reglucosylation-refolding cycle and the simpler glycoprotein degradation pathway in yeast. Here, we engineered the glycoprotein QC system of Kluyveromyces marxianus by introducing key human components and modifying native pathways. Human UGGT1 or UGGT2 enhanced soluble and secretory glycoprotein production in an activity-dependent manner, with further improvements achieved by co-expressing the human cochaperone SEP15 and reducing native glucosidase II trimming. Human EDEM2 delayed endoplasmic reticulum-associated degradation and increased secretion. Combining these strategies enhanced the production of diverse N-glycoproteins, including Fc, γ-glutamyl hydrolase, fungal xylanase, and Fc-fusion therapeutics, by up to ∼12-fold, demonstrating an effective strategy for engineering human-like glycoprotein QC in yeast to improve glycoprotein production.

Flavonolignans are pharmacologically important plant natural products whose production has been limited to extraction from natural sources. Their defining transformation, the oxidative coupling of flavonoid and phenylpropanoid precursors, was demonstrated to be enzyme-catalyzed. Here we show that milk thistle flavonolignans can also be assembled through an abiotic, light-driven process in vitro. Photocatalytic coupling of taxifolin and coniferyl alcohol generated the major natural flavonolignans, including silychristins, whose synthesis from native precursors has not previously been reproduced in vitro. Importantly, we identify environmental metal ions as programmable regulators of reaction outcome. Fe[3+] and Al[3+] selectively direct radical coupling toward silychristin formation, whereas Mn[2+] primarily enhances catalytic efficiency. Mechanistic analyses indicate that metal coordination of flavonoid substrates governs regioselective radical attack and molecular architecture of final products. Leveraging this principle, we establish a metal-programmable photocatalytic platform for the synthesis of both natural and new-to-nature flavonolignans. Integration of this abiotic chemistry with engineered Saccharomyces cerevisiae enabled de novo flavonolignan production directly from sugar in a one-pot photo-biohybrid process for the first time, establishing a versatile strategy for the sustainable production and diversification of flavonolignans.

Two-stage bioprocesses which decouple cell growth from product synthesis are an attractive approach to biomanufacturing. However high levels of production in stationary phase cultures often suffer from a progressive decline in metabolism. We demonstrate that in E. coli pyruvate accumulation, an inevitable consequence of high-flux metabolism, acts as a major inhibitor of stationary-phase glucose uptake. To address this limitation, we introduce a redesigned central metabolic architecture, the gluconate-bypass (GBP), which reroutes carbon flux around glucose-6-phosphate to sustain metabolic activity during stationary phase production. This architecture provides two key advantages: it decouples glucose uptake from pyruvate mediated inhibition, enabling prolonged stationary phase productivity, and glucose oxidation intrinsically co-generates the reducing cofactor NADPH to support biosynthetic pathways that require NADPH. We validated this architecture using the NADPH dependent production of L-alanine as a representative case study. Implementation of the GBP metabolism generated a self regulating host that achieved a record alanine titer of 197 g L[-1] and extended production longevity by 1.6 fold, resulting in an improved production yield of 94%. Together, these results demonstrate that the GBP metabolism supports robust stationary phase biosynthesis and provides a versatile framework for efficient production of pyruvate derived chemicals.

Microbial coculture can integrate advantages and overcome the metabolic imbalance of individual species. Programming strain interactions represents a common routine for synthetic microbial communities with distinct species, which causes difficulties and redundant workloads in interaction construction before being available as chassis hosts. This study explores yeast-bacterium consortium without engineered interactions for the co-inducible relayed synthesis of natural products. The Komagataella phaffii-Escherichia coli consortium is explored for co-growth under selected conditions. Low-level glucose- and blue light-responsive transcriptional systems are rebuilt separately for each host, allowing single-signal co-induced activation of compound synthesis in coculture. Pathway redirection, genome mining, and rewiring of key targets for acyl donor degradation result in efficient production of the reporter molecule simvastatin (26.2 mg l[-1]) through living consortium cultured on simple carbon source. Inducible biosynthesis of another reporter compound (2S)-naringenin (165.6 mg l[-1]) further validates the extendibility of this community. The described platform represents a breakthrough in engineering microbial consortium for biosynthesis.

Efficient microbial conversion of heterogeneous carbohydrate mixtures remains a central bottleneck in industrial biotechnology, yet scalable chassis architectures capable of coordinated multi-sugar utilization are limited. Recently, substrate-informed metabolic alignment enabled balanced glucose-xylose co-utilization in Corynebacterium glutamicum. Here, we examine whether this alignment principle is scalable and transferable to complex industrial feedstocks. We progressively expanded the substrate spectrum of C. glutamicum through genome-encoded integration of mannose, xylose, arabinose, galactose, and rhamnose utilization modules, establishing a stable seven-sugar chassis. Rather than relying on adaptive evolution or high-copy expression, pathway capacities were quantitatively aligned with native uptake and central metabolism. Progressive expansion increased total volumetric carbon uptake and attenuated substrate hierarchy without compromising transcriptional stability. Complementary 13C tracer experiments showed that carbon from all seven sugars was incorporated into biomass in proportion to substrate utilization while revealing characteristic precursor-level incorporation patterns consistent with their distinct metabolic entry routes. Using authentic spent sulfite liquor (SSL) as a chemically heterogeneous validation substrate, the engineered strain achieved near-complete depletion of all fermentable sugars and substantially accelerated conversion kinetics compared to the wild type. Transfer of the aligned substrate module into a glucose-optimized glutarate production strain enabled sustained fed-batch conversion of SSL to 30 g L-1 glutarate without pathway erosion or performance decline. Together, these results demonstrate that substrate-informed metabolic engineering can provide a scalable and industrially transferable strategy for chassis development and position C. glutamicum as a robust multi-substrate platform for heterogeneous renewable feedstocks.

In the model bacterium Escherichia coli, the Embden-Meyerhof-Parnas (EMP) glycolysis pathway is the primary route for carbohydrate metabolism. However, alternative metabolic routes can be activated depending on genetic configurations and available carbon sources. In this study, E. coli strains lacking key enzymes of the lower EMP pathway-phosphoglycerate kinase or enolase-were subjected to long-term adaptive evolution in continuous culture under a medium swap regime, for growth on D-xylose as the sole carbon and energy source. Through metabolic [13]C-labeling experiments on evolved isolates, we found that carbon flux was predominantly rerouted via the Entner-Doudoroff pathway, known to remain silent when wild type cells are fed with D-xylose. To investigate the molecular basis of this growth adaptation, we identified the mutations fixed in the genomes of evolved prototrophic isolates and conducted comprehensive transcriptomic and proteomic analyses. Our study revealed that mutations in key enzymes and transcriptional regulators at various metabolic branching points were essential for effective growth on D-xylose, enabling optimized partitioning of carbon through central metabolism despite the loss of the canonical EMP glycolytic route. The multi-layered regulatory and metabolic adaptations identified in the evolved strains demonstrate the complex nature of evolutionary trajectories and underscore the potential of adaptive evolution to optimize metabolic network function and enzyme utilization in ways that extend rational engineering approaches.

Perfusion Chinese hamster ovary (CHO) cell culture offers potential benefits over fed-batch cultures, including higher productivity and a reduced production footprint. However, maintaining a stable profile of product quality attributes (PQAs) over the entirety of a perfusion culture has proven to be a challenge across the biopharmaceutical industry, which is critical for ensuring lot-to-lot consistency. A deeper understanding of the underlying molecular mechanisms of PQA variability and productivity can allow for more targeted interventions. In this study, [13]C metabolic flux analysis ([13]C MFA) was performed on perfusion bioreactors at both early and late stage timepoints to identify potential metabolic bottlenecks. Additionally, RNA-level expression of transporters and enzymes throughout metabolism was measured using RNAseq analysis. Central carbon metabolism in late stage culture was found to have significantly higher flux throughout glycolysis and the tricarboxylic acid (TCA) cycle. In nucleotide sugar synthesis pathways, the ratio of GDP-Fucose synthesis flux to GDP-Mannose synthesis flux was reduced at later cell culture stages, indicating a potential bottleneck for maintaining stable profiles of fucosylated and mannosylated glycans. This bottleneck could potentially be explained by changes in expression of the enzymes in these synthesis pathways as well as a limited pool of NADPH. This work represents the first report demonstrating the application of [13]C MFA in early and late-stage perfusion cultures, elucidating potential molecular bottlenecks contributing to unstable PQA profiles.

Eupatilin (EUP) and jaceosidin (JAC) are bioactive polymethoxylated flavones (PMFs) derived from Artemisia argyi. However, their sustainable production is hindered by the low abundance of this plant and the challenges of regioselective chemical synthesis. One key obstacle in microbial PMF biosynthesis lies in coordinating precursor supply with P450-dependent oxidation and SAM-dependent O-methylation. In this study, we report the first de novo biosynthesis of EUP and JAC in Saccharomyces cerevisiae using a modular "backbone-first, tailoring-next" strategy. Here, a tyrosine-to-(2S)-naringenin module was coupled with a CYP450 tailoring cascade (FNSII-F3'H-F6H) to produce 6-hydroxyluteolin, followed by product-specific O-methylation via complementary flavonoid O-methyltransferases. Through enzyme screening, precursor-flux optimization, gene-dosage tuning, and NADPH/SAM cofactor engineering, the optimized strains achieved de novo production titers of 27.3 mg/L EUP and 42.2 mg/L JAC. Further optimization of pH and staged-feeding in a 5-L bioreactor increased the mean peak titers to 70.5 mg/L EUP and 63.3 mg/L JAC at 120 h. This work establishes a yeast platform for PMF biosynthesis and provides a general framework for engineering P450- and OMT-dominated flavonoid pathways.

Lignin-rich industrial streams represent an abundant but underutilized source of renewable aromatic carbon. Efficient biological conversion requires microbial hosts capable of metabolizing chemically diverse lignin-derived aromatic compounds; however, such capabilities are typically found in environmental bacteria, which are constrained by physiological and metabolic limitations. Sphingobium lignivorans SYK-6 harbors extensive aromatic catabolic pathways, but its inability to utilize glucose and its methionine auxotrophy have limited its use as a production host. Here, we identified the metabolic basis of these constraints and systematically rewired the underlying pathways to overcome them. Introduction of a heterologous glucose transporter, reconstruction of methionine biosynthesis, chromosomal integration of pathway genes, and adaptive laboratory evolution collectively enabled robust growth on glucose while eliminating methionine auxotrophy. The engineered strain converted lignin-derived aromatics in oxygen-soda-anthraquinone pulping black liquor derived from Japanese cedar, achieving high-yield production of the polymer precursor 2-pyrone-4,6-dicarboxylic acid (PDC) (2.71 g/L and >130 mol% conversion relative to major quantified aromatics). We further show that gluconolactonase can substitute for 6-phosphogluconolactonase in the Entner-Doudoroff pathway, demonstrating that central carbon metabolism can accommodate functionally analogous enzymes. Together, these results provide a metabolically rewired SYK-6 strain as a platform for the valorization of industrial lignin streams and suggest that overcoming physiological and metabolic constraints can enable non-model aromatic-degrading bacteria to function as industrial production hosts.

Biopolymer chitin has properties such as biocompatibility, strength, chemical functionality, and biodegradability that are attractive for applications. It could be biotechnologically produced using fungi, but the economical feasibility depends on the chitin content in cells. However, chitin content in cells is governed by several cellular regulatory pathways and has complex genetic underpinnings. Here, adaptive laboratory evolution (ALE) in presence of caspofungin (CAS) was successfully devised for selecting chitin overaccumulation in Saccharomyces cerevisiae. After ∼200 generations of independent asexual adaptive evolution of S. cerevisiae lineages originating from wild type (BY4742), kre6Δ0 strain and an UV treated population of wild type cells, all lineages tolerated the highest introduced concentration of CAS and most of them overproduced chitin. The highest specific chitin content was found in the ALE mid-point population of a wild type -based lineage. Whole genome sequencing of parental and evolved populations revealed 88 unique single-nucleotide variants affecting 43 coding sequences. High-frequency variants detected in the evolved population with the highest chitin content (i.e., Fks1 p. Pro647Arg and p. Arg1357Cys, Fks2 p. Arg1203∗, and Ecm21 p. Glu642∗) were re-engineered into the wild type strain. The re-engineering revealed that chitin content in cells could be increased up to 7-fold with a specific variant combination (i.e., Fks1 p. Pro647Arg, Fks2 p. Arg1203∗, and Ecm21 p. Glu642∗). Interestingly, they did not improve CAS tolerance. Biotechnological production requires industrial compatibility which has been proven for S. cerevisiae. The variants identified here form a basis for developing an industrial S. cerevisiae strain for biotechnological production of animal-free chitin.

The sexual life cycle represents a natural engine for promoting genetic diversity in yeast. However, harnessing it for controllable, rapid evolution in industrial yeast strains remains a big challenge. Here, we developed quantitative mating and sporulation methods for an industrial yeast Komagataella phaffii. Functional genes for its sexual life cycle were identified, and deficiency of key sporulation targets generated stable diploids with enhanced stress tolerance and recombinant protein production. A fully programmable life-cycle system was constructed, for which mating and sporulation are orthogonally controlled by different inducible signals. Leveraging this system, we developed an in situ mating-sporulation shift (iMASS) platform in liquid culture. Applications of iMASS under diverse selection pressures accelerated cell evolution, generating strains with improved robustness and protein productions. Our work proposes a paradigm that integrates rational design with controlled sexual reproduction for directed microbial evolution.