Metabolic Engineering

Eukaryotic cells compartmentalize metabolic pathways in organelles to achieve optimal reaction conditions and avoid crosstalk with other factors in the cytosol. Increasingly, engineers are researching ways in which synthetic compartmentalization could be used to address challenges in metabolic engineering. Here, we identified that norcoclaurine synthase (NCS), the enzyme which catalyzes the first committed reaction in benzylisoquinoline alkaloid (BIA) biosynthesis, is toxic when expressed cytosolically in Saccharomyces cerevisiae and, consequently, restricts (S)-reticuline production. We developed a compartmentalization strategy that alleviates NCS toxicity while promoting increased (S)-reticuline titer, achieved through efficient targeting of toxic NCS to the peroxisome while, crucially, taking advantage of the free flow of metabolite substrates and product across the peroxisome membrane. We identified that peroxisome protein capacity in S. cerevisiae becomes a limiting factor for further improvement of BIA production and demonstrate that expression of engineered transcription factors can mimic the oleate response for larger peroxisomes, further increasing BIA titer without the requirement for peroxisome induction with fatty acids. This work specifically addresses the challenges associated with toxic NCS expression and, more broadly, highlights the potential for engineering organelles with desired characteristics for metabolic engineering.

To comparatively evaluate cellular constraints on recombinant monoclonal antibody (mAb) production by Chinese Hamster Ovary (CHO) cells, we analysed the transcriptomes of 24 clonally derived CHO cell lines engineered with PiggyBac transposon technology to stably produce four recombinant monoclonal antibodies (mAbs) at varying specific production rates. Fed-batch cultures were sampled at exponential (day 5) and stationary (day 10) phases of culture for analysis by RNA-Seq. Recombinant mRNAs accounted for a large proportion of total mRNA across all clones, and efficient use of heavy chain (HC) mRNA to synthesise recombinant mAb (qP per HC mRNA) varied significantly with respect to both mAb product and cell line. Comparative bioinformatic analyses of CHO transcriptomes focussed on mAb specific production rate and utilised both data-driven and hypothesis-led approaches, specifically (i) production or non-production of recombinant mAb, (ii) changes in the abundance of functional groups of mRNAs abundance with mAb specific production rate and (iii) comparative analysis of informatically-mined gene subsets associated with cellular functions hypothesised to impact recombinant mAb synthesis and secretion. These analyses revealed widespread constitutive and adaptive changes in mRNA abundance associated with mAb production across a variety of cellular functions. Typically, most mechanistically consistent changes in mRNA abundance co-varying with mAb production were evident at the stationary phase sample point. These data revealed both recombinant mAb-specific limitations on cellular synthetic capacity and a generic adaptive strategy used by CHO cells to support high-level mAb production. The latter was achieved by directed and permissive regulation of endoplasmic reticulum and other processes to accommodate increased synthetic flux.

13C-metabolic flux analysis (13C-MFA) is a crucial technique that experimentally determines metabolic flux distribution. Although precision of each flux strongly depends on tracer labeling pattern, its optimization remains challenging. We developed an integrated platform, OpenMebius2, a graphical user interface (GUI)-based software for 13C-MFA that includes a tracer labeling pattern suggestion function to support subsequent experiments. The proposed function leverages metabolic flux distributions and their 95 % confidence intervals obtained using low-cost 13C-labeled substrates to evaluate hypothetical parallel labeling scenarios and predict improvements in flux estimation precision. Availability and implementationThis software runs on Linux, macOS, and Windows. The source code and binary files are available at https://github.com/metabolic-engineering/OpenMebius2 under the PolyForm Noncommercial License 1.0.0.

BackgroundIn response to rising CO2 emissions driving global warming, there is an urgent need for a transition toward a sustainable bioeconomy. Photo-biotechnological processes based on oxygenic photosynthesis hold high potential for achieving CO2 neutrality and in this regard, cyanobacteria have emerged as promising biocatalysts. Rational metabolic engineering of cyanobacteria depends on a thorough understanding of native regulatory mechanisms governing primary metabolism, which can limit metabolic flux through specific pathways and, consequently, the formation of target products. Recent insights have identified a key regulatory node at the 2,3-bisphosphogylcerate-independent phosphoglycerate mutase (PGAM) reaction, where the metabolic flux from newly fixed carbon is redirected from the Calvin-Benson-Bassham (CBB) cycle towards lower glycolysis. This metabolic valve is controlled by the small inhibitor protein PirC, whose binding to PGAM is determined by the central signal transduction protein PII. ResultsIn this study, we exploit the PirC-PGAM interaction as a novel target for regulatory metabolic engineering in the model cyanobacterium Synechocystis sp. PCC 6803 (Synechocystis). Chassis strains with engineered control of PGAM, defined as PGAM-ON or PGAM-OFF states, were generated using two complementary approaches: tuning pgam gene expression and modulating PirC abundance to regulate PGAM activity. The effectiveness of this regulatory engineering strategy was demonstrated by redirecting carbon flux toward two representative, naturally occurring products: sucrose, produced via gluconeogenesis fueled by the Calvin-Benson-Bassham (CBB) cycle, and succinate, an intermediate of the tricarboxylic acid (TCA) cycle. Narrowing the PGAM valve resulted in a threefold increase in sucrose accumulation. In contrast, opening the PGAM valve by relieving PGAM inhibition through pirC deletion or separate pgam overexpression resulted in up to an 18-fold increase in succinate excretion. Furthermore, similar genetic configurations were applied to enhance production of a heterologous compound, isoprene, derived from pyruvate. ConclusionsThis study establishes the PGAM valve as a tunable control point for the rational re-direction of carbon flux in Synechocystis and highlights small regulatory proteins as powerful targets for metabolic engineering. Together, these findings provide proof of concept for an advanced level of molecular engineering in cyanobacteria and to fully harness their biocatalytic potential in future photosynthesis-driven biotechnological applications.

Microbial fermentation for chemical production is becoming more broadly adopted as an alternative to petrochemical refining. Fermentation typically relies on sugar as a feedstock, however, one-carbon compounds like methanol are an attractive alternative as they can be derived from organic waste and natural gas. This study focused on engineering methanol assimilation in the yeast Saccharomyces cerevisiae. Three methanol assimilation pathways were engineered and tested: a synthetic xylulose monophosphate (XuMP), a hybrid methanol dehydrogenase-XuMP, and a bacterial ribulose monophosphate (RuMP) pathway, with the latter identified as the most effective at assimilating methanol. Additionally, 13C-methanol tracer analysis uncovered a native capacity for methanol assimilation in S. cerevisiae, which was optimized using Adaptive Laboratory Evolution. Three independent lineages selected in liquid methanol-yeast extract medium evolved premature stop codons in YGR067C, which encodes an uncharacterised protein that has a predicted DNA-binding domain with homology to the ADR1 transcriptional regulator. Adr1p regulates genes involved in ethanol metabolism and peroxisomal proliferation, suggesting YGR067C has a related function. When one of the evolved YGR067C mutations was reverse engineered into the parental CEN.PK113-5D strain, there were up to 5-fold increases in 13C-labelling of intracellular metabolites from 13C-labelled methanol when 0.1 % yeast extract was a co-substrate, and a 44 % increase in final biomass. Transcriptomics and proteomics revealed that the reconstructed YGR067C mutation results in down-regulation of genes in the TCA cycle, glyoxylate cycle, and gluconeogenesis, which would normally be up-regulated during growth on a non-fermentable carbon source. Combining the synthetic RuMP and XuMP pathways with the reconstructed Ygr067cp truncation led to further improvements in growth. These results identify a latent methylotrophic metabolism in S. cerevisiae and pave the way for further development of native and synthetic one-carbon assimilation pathways in this model eukaryote.

The Warburg effect, the preferential conversion of glucose-derived pyruvate to lactate despite available oxygen, is a key feature of Chinese hamster ovary (CHO) cell culture. Lactate accumulation in recombinant protein-producing cell culture is an inefficient usage of glucose, as well as being deleterious to cells. Lactate accumulation lowers culture pH, requiring base addition to maintain bioreactor pH setpoint, which subsequently leads to hyperosmolarity, adversely impacting cell growth, productivity and product quality. A key driver for the Warburg effect, and hence lactate accumulation, is the need to regenerate NAD+ consumed during glycolysis. Since oxidative phosphorylation (OXPHOS) has limited capacity to recycle NADH back to NAD+ at high glycolytic fluxes, cells rely on lactate dehydrogenase (LDH) to convert pyruvate to lactate, simultaneously regenerating NAD+ and sustaining glycolysis. Thus, providing the cells capacity to generate more NAD+ would decrease the reliance on the Warburg effect. In this study, feeding the NAD+ precursor nicotinamide (NAM) leads to reversal of the Warburg effect, inducing the "lactate shift" three days earlier in cell culture and reducing peak lactate concentration by 40%. Transcriptomic analysis further confirms this metabolic shift, with an upregulation of key mitochondrial electron transport chain genes. These results identify NAD+/NADH balance as a key regulator of the Warburg effect and demonstrate NAM supplementation as a simple, cost-effective strategy to mitigate lactate accumulation and improve metabolic efficiency in CHO cell cultures.

While photosynthetic cyanobacteria are potential biotechnological hosts for light-driven production of sustainable chemicals from CO2, engineering more efficient strains is critical for the development of competitive industrial processes. This study demonstrates significantly enhanced production of the soluble bioplastic precursor (R)-3-hydroxybutyrate (3HB) that has been engineered based on the polyhydroxybutyrate (PHB) pathway in photoautotrophic cyanobacterium Synechocystis sp PCC 6803. As the key novelty, we generated a library of engineered 3HB pathway variants that express the three key heterologous pathway enzymes PhaA, PhaB and TesB at varying efficiencies, followed by the screening of most efficient 3HB producers. This was achieved by placing each of the pathway enzymes under the translational regulation of three alternative RBSs in different combinations, resulting in strains with wide dynamic range of 3HB productivities. The best strains accumulated over 5 gl-1 under 200 mol photons m-2s-1 and 3% CO2 in a 14-day flask batch culture, with the highest titer reaching 12 gl-1, corresponding to nearly 3 gl-1 d-1 during the peak production phase. These are the highest 3HB production levels reported so far in cyanobacteria, and comparable to those previously established in heterotrophic production systems. Proteomic comparison of selected strains revealed that the different RBS combinations result in varying expression patterns of the pathway proteins, and that the strain-specific enzyme levels remained relatively constant over the monitored six-day period. The results show that altering the levels of the target pathway enzymes can dramatically improve product yield in Synechocystis, while even very small quantitative differences in the strain-specific expression profiles can have marked effects on the production efficiency. This could be a general tool for optimizing engineered pathways in cyanobacteria, provided that the flux to the end-product is not critically restricted by substrate availability but determined by the balance between the consecutive pathway steps.

Brain and neurological diseases are influencing more than one billion worlds people. Nervonic acid (cis-15-tetracosenoic acid, C24:1 {Delta}15) benefits the treatment of neurological diseases and the health of brain. Currently, the sources of nervonic acid are limited to the seeds of a couple of plants. In this study, we employed the oleaginous yeast Yarrowia lipolytica to overproduce nervonic acid oil by systematic metabolic engineering. First, engineering the fatty acid elongation (FAE) pathway by expressing a heterologous {beta}-ketoacyl-CoA synthase gene CgKCS enabled the production of nervonic acid in Y. lipolytica. Second, modulation of endogenous pathways by expressing a C16:0-acyl-CoA preferred fatty acid elongase gELOVL6 together with a C18:0-acyl-CoA preferred fatty acid desaturase MaOLE2 increased the content of nervonic acid in total fatty acids (TFA). Third, iterative expression of CgKCS, gELOVL6 and MaOLE2 at the genomic loci of rDNA, FAD2, TGL4, GSY1 and SNF1 dramatically improved the production of nervonic acid. Fourth, the biosynthesis of both nervonic acid and lipids were further enhanced by expression of the MaOLE2-CgKCS fusion protein and glycerol-3-phosphate acyltransferases (GPAT) and diacylglycerol acyltransferases (DGAT) from Malania oleifera in the endoplasmic reticulum (ER) membrane. Fifth, an ER structure regulator YlINO2 was identified in Y. lipolytica and the overexpression of YlINO2 led to a 39.3% increase in lipid production. Next, pilot-scale fermentation in 50-L reactor using the strain YLNA9 exhibited a lipid titer of 96.7 g/L and a nervonic acid titer of 17.3 g/L, the highest reported titer to date for de novo nervonic acid production. We also found that disruption of the AMP-activated S/T protein kinase SNF1 increased the ratio of nervonic acid (C24:1) to lignoceric acid (C24:0) by 61.6% and a ratio of 3.5:1 (nervonic acid to lignoceric acid) was achieved in the strain YLNA10. Finally, a proof-of-concept purification and separation of nervonic acid were performed and the purity of it reached 98.7%. This study suggested that oleaginous yeasts are attractive hosts for the cost-efficient production of nervonic acid and possibly other very long-chain fatty acids (VLCFAs).

Short-chain esters are versatile chemicals with use as flavors, fragrances, solvents, and fuels. The de novo ester biosynthesis consists of diverging and converging pathway submodules, which is challenging to engineer to achieve optimal metabolic fluxes and selective product synthesis. Compartmentalizing the pathway submodules into specialist cells that facilitate pathway modularization and labor division can present a promising solution. Here, we engineered a synthetic Escherichia coli coculture with the compartmentalized sugar utilization and ester biosynthesis pathways to produce isobutyl butyrate from a mixture of glucose and xylose. To compartmentalize the sugar-utilizing pathway submodules, we engineered a xylose-utilizing E. coli specialist that selectively consumes xylose over glucose and bypasses the carbon catabolite repression (CCR) while leveraging the native CCR machinery to activate a glucose-utilizing E. coli specialist. Upon compartmentalizing the isobutyl butyrate pathway submodules into these sugar-utilizing specialist cells, a robust synthetic coculture could be engineered to selectively produce isobutyl butyrate at a level of 392 mg/L, about 31-fold higher than the monoculture.

Cell-free protein expression has become a widely used research tool in systems and synthetic biology and a promising technology for protein biomanufacturing. Cell-free protein synthesis relies on in-vitro transcription and translation processes to produce a protein of interest. However, transcription and translation depend upon the operation of complex metabolic pathways for precursor and energy regeneration. Toward understanding the role of metabolism in a cell-free system, we developed a dynamic constraint-based simulation of protein production in the myTXTL E. coli cell-free system with and without electron transport chain inhibitors. Time-resolved absolute metabolite measurements for [M] = 63 metabolites, along with absolute concentration measurements of the mRNA and protein abundance and measurements of enzyme activity, were integrated with kinetic and enzyme abundance information to simulate the time evolution of metabolic flux and protein production with and without inhibitors. The metabolic flux distribution estimated by the model, along with the experimental metabolite and enzyme activity data, suggested that the myTXTL cell-free system has an active central carbon metabolism with glutamate powering the TCA cycle. Further, the electron transport chain inhibitor studies suggested the presence of oxidative phosphorylation activity in the myTXTL cell-free system; the oxidative phosphorylation inhibitors provided biochemical evidence that myTXTL relied, at least partially, on oxidative phosphorylation to generate the energy required to sustain transcription and translation for a 16-hour batch reaction.

Lactones constitute a family of aroma compounds found in fruits, flowers and vegetables and are in high demand in the food industry. Previous studies have reported biotransformation of castor beans-extracted ricinoleic acid to {gamma}-decalactone using oleaginous yeast Yarrowia lipolytica. Given the potential toxicities associated with castor beans, we have used a metabolic flux-engineering approach to produce {gamma}-decalactone from oleic acid in Saccharomyces cerevisiae. Intracellular conversion of oleic acid to ricinoleic acid was achieved by the expression of oleate hydroxylase Fah12 from ergot fungus Claviceps purpurea. Glycerol-3-phosphate dehydrogenase-mediated glycerol synthesis was identified as the major metabolic diversion of oleic acid that negatively impacts {gamma}-decalactone yields. Chemogenomic profiling analysis revealed that the tryptophan biosynthetic pathway provides resistance to {gamma}-decalactone-mediated toxicity in yeast. Overexpression of tryptophan transporter Tat1 enhanced {gamma}-decalactone production by about 3- fold. Deficiency of genes encoding the cytoplasmic fatty acyl CoA synthetases FAA1 or FAA4 alone did not significantly influence {gamma}-decalactone production. However, deficiency of peroxisomal FAA2 drastically diminished the yield of {gamma}-decalactone. Thus, this study uncovers the metabolic barriers to oleic acid-to-{gamma}-decalactone conversion and identifies Faa2 as an essential element in this biotransformation.

The photosynthetic cyanobacterium Synechocystis sp. PCC 6803 offers a promising sustainable solution for simultaneous CO2 fixation and compound bioproduction. While various heterologous products have now been synthesised in Synechocystis, limited genetic tools hinder further strain engineering for efficient production. Here, we present a versatile CRISPR activation (CRISPRa) system for Synechocystis, enabling robust multiplexed activation of both heterologous and endogenous targets. Following tool characterisation, we applied CRISPRa to explore targets influencing biofuel production, specifically isobutanol (IB) and 3-methyl-1-butanol (3M1B), demonstrating a proof-of-concept approach to identify key reactions constraining compound biosynthesis. Notably, individual upregulation of target genes, such as pyk1, resulted in up to 4-fold increase in IB/3M1B formation while synergetic effects from multiplexed targeting further enhanced compound production, highlighting the value of this tool for rapid metabolic mapping. Interestingly, activation efficacy did not consistently predict increases in compound formation, suggesting complex regulatory interactions influencing bioproduction. This work establishes the first CRISPRa system in cyanobacteria, providing an adaptable platform for high-throughput screening, metabolic pathway optimisation and functional genomics. Our CRISPRa system provides a crucial advance in the genetic toolbox available for Synechocystis and will facilitate innovative applications in both fundamental research and metabolic engineering in cyanobacteria.

Cofactor imbalance obstructs the productivities of metabolically engineered cells. Herein, we employed a minimally perturbing system, xylose reductase and lactose (XR/lactose), to increase levels of a pool of sugar-phosphates which are connected to the biosynthesis of NAD(P)H, FAD, FMN and ATP in Escherichia coli. The XR/lactose system could increase the amounts of the precursors of these cofactors and was tested with three different metabolically engineered cell systems (fatty alcohol biosynthesis, bioluminescence light generation and alkane biosynthesis) with different cofactor demands. Productivities of these cells were increased 2-4-fold by the XR/lactose system. Untargeted metabolomic analysis revealed different metabolite patterns among these cells; demonstrating that only metabolites involved in relevant cofactor biosynthesis were altered. The results were also confirmed by transcriptomic analysis. Another sugar reducing system (glucose dehydrogenase, GDH) could also be used to increase fatty alcohol production but resulted in less yield enhancement than XR. This work demonstrates that the approach of increasing cellular sugar phosphates can be a generic tool to increase in vivo cofactor generation upon cellular demand for synthetic biology. TeaserUse of sugar and sugar reductase to increase sugar phosphates for enhancing in situ synthesis of cofactors upon cellular demand for synthetic biology.

Converting industrial side streams into value-added chemicals using microbial cell factories is of increasing interest, as such processes offer solutions to reduce waste and production costs. However, developing new, efficient cell factories for precision fermentation remains challenging due to limited knowledge about their metabolic capabilities. Here, we investigate the lactose and galactose metabolism of the non-conventional yeast Sungouiella intermedia (formerly Candida intermedia), using knowledge-matching of high-quality genome-scale metabolic model (GEM) with extensive experimental analysis and determine its potential as a future cell factory on lactose-rich industrial side-streams. We show that this yeast possesses the conserved Leloir pathway as well as an oxidoreductive galactose catabolic route. Contextualization of RNAseq data into Sint-GEM highlights the regulatory mechanisms on the oxidoreductive pathway and how this pathway can enable adaptation to diverse environments. Model simulations, together with experimental data from continuous and batch bioreactors, indicate that S. intermedia uses upstream enzymes of the oxidoreductive pathway, in a condition-dependent manner, and produce the sugar alcohol galactitol as a carbon overflow metabolite, coupled to redox co-factor balancing during both lactose and galactose growth. Furthermore, the new metabolic insights facilitated the development of an improved bioprocess design, where an engineered S. intermedia strain could achieve galactitol yields of >90% of the theoretical maximum at improved production rates using the industrial side-stream cheese whey permeate as feedstock. Additional strain engineering resulted in galactitol-to-tagatose conversion, proving the versatility of the future production host. Overall, this work sheds new light on the intrinsic interplay between parallel metabolic pathways that shape the lactose and galactose catabolism in S. intermedia. It also demonstrates how a GEM combined with experimental analysis can work in synergy to fast-forward metabolic characterization and development of new, non-conventional yeast cell factories. HighlightsO_LIAn oxidoreductive pathway functions in concert with the Leloir pathway for galactose catabolism. C_LIO_LIGEM predicts that galactitol secretion enables efficient carbon overflow metabolism and maintains redox balance. C_LIO_LIKnowledge-matching of GEM with experimental results highlights cell factory potential. C_LIO_LIHigh galactitol yields and proof-of-concept tagatose production using whey permeate as feedstock. C_LI

We demonstrate the use of two-stage dynamic metabolic control to manipulate feedback regulation in central metabolism and improve stationary phase biosynthesis in engineered E. coli. Specifically, we report the impact of dynamic control over two enzymes: citrate synthase, and glucose-6-phosphate dehydrogenase, on stationary phase fluxes. Firstly, reduced citrate synthase levels lead to a reduction in -ketoglutarate, which is an inhibitor of sugar transport, resulting in increased stationary phase glucose uptake and glycolytic fluxes. Reduced glucose-6-phosphate dehydrogenase activity activates the SoxRS regulon and expression of pyruvate-ferredoxin oxidoreductase, which is in turn responsible for large increases in acetyl-CoA production. The combined reduction in citrate synthase and glucose-6-phosphate dehydrogenase, leads to greatly enhanced stationary phase metabolism and the improved production of citramalic acid enabling titers of 126{+/-}7g/L. These results identify pyruvate oxidation via the pyruvate-ferredoxin oxidoreductase as a "central" metabolic pathway in stationary phase E. coli, which coupled with ferredoxin reductase comprise a pathway whose physiologic role is maintaining NADPH levels. HighlightsO_LIDynamic reduction in -keto-glutarate pools alleviate inhibition of PTS dependent transport improving stationary phase sugar uptake. C_LIO_LIDynamic reduction in glucose-6-phosphate dehydrogenase activates pyruvate flavodoxin/ferredoxin oxidoreductase and improves stationary acetyl-CoA flux. C_LIO_LIPyruvate flavodoxin/ferredoxin oxidoreductase is responsible for large stationary phase acetyl-CoA fluxes under aerobic conditions. C_LIO_LIProduction of citramalate to titers 126 {+/-} 7g/L at > 90 % of theoretical yield. C_LI

CO2 recycle is crucial to the global carbon neutrality. Though cyanobacteria are known to be photoautotrophic cell factories capable of converting CO2 into valuable chemicals, their slower growth rate and lower biomass accumulation compared to those of the heterotrophic organisms significantly restrict their application at commercial scale. The newly discovered marine cyanobacterium, Synechococcus sp. PCC 11901 (hereafter PCC 11901) offers several advantages like rapid growth, high biomass and high salinity tolerance, and could become a new generation of cyanobacterial chassis. To promote its application, in this study we developed genetic toolboxes applicable to PCC11901. First, a cobalamin (VB12)-independent chassis was constructed, allowing for its cheaper cultivation. Second, genome copy numbers and transformation methods were respectively measured and optimized. The 14 neutral sites were identified and characterized within the genome PCC 11901, providing locations for genetic integration of exogenous cassettes. Subsequently, libraries were developed, reaching an expression range of approximately 800 folds for constitutive promoters and an induction fold of up to approximately 400 for inducible promotor, respectively. As a proof of concept of its utilization, we engineered the synthetic pathways of glucosylglycerol (GG) into PCC 11901 using the established toolboxes, yielding 590.41 {+/-} 21.48 mg/L for GG production. Notably, we found the cobalamin-independent PCC 11901 chassis exhibited superior self-sedimentation ability compared to the wild-type chassis. Our work here made it possible to develop the fast-growing PCC 11901 as efficient carbon-neutral cell factory in the future.

Hemicelluloses are important dietary fibers and a key component of lignocellulosic biomass. Despite numerous observations for fluorescently tagged cellulose synthases, the subcellular journeys and biochemical activities of intracellular cellulose synthase-like enzymes such as {beta}-mannan synthases (ManS) remain largely unexplored. This study identifies C-terminal fluorescent protein tags that maintain ManS activity in the yeast to accelerate the Design, Build, Test, Learn cycles for polysaccharide biosynthesis. Using the Amorphophallus konjac ManS as a case study, we demonstrate that the enzyme co-localizes with a known yeast marker for the Golgi apparatus despite the toxic effects of plant glucomannan accumulation in Pichia pastoris. The ManS first transmembrane domain was found to be critical for the punctate localization of the enzyme, its overall expression level and its function. Additionally, we explored how fluorescently tagged ManS is influenced by genetic or chemical perturbations of native yeast cell wall components, such as reducing protein mannosylation and severely disrupting {beta}-1,3-glucans. Finally, we identified alternative feeding strategies and episomal vectors for Pichia, which were extended to Saccharomyces cerevisiae, to accelerate hemicellulose research. We propose that expanding the Plant MoClo-compatible plasmid repertoire is essential to swiftly prototype carbohydrate-active enzymes in yeast before proceeding with more time-intensive analyses in plants. Requiring only hours or days instead of weeks or months for plant transformation/regeneration, our yeast prototyping strategies can de-risk the bioengineering of carbohydrate-active enzymes.

Formate is a promising energy carrier that could be used to transport renewable electricity. Some acetogenic bacteria, such as Eubacterium limosum, have the native ability to utilise formate as a sole substrate for growth, which has sparked interest in the biotechnology industry. However, formatotrophic metabolism in acetogens is poorly understood, and a systems-level characterization in continuous cultures is yet to be reported. Here we present the first steady-state dataset for E. limosum formatotrophic growth. At a defined dilution rate of 0.4 d-1, there was a high specific uptake rate of formate (280{+/-}56 mmol/gDCW/d), however, most carbon went to CO2 (150{+/-}11 mmol/gDCW/d). Compared to methylotrophic growth, protein differential expression data and intracellular metabolomics revealed several key features of formate metabolism. Upregulation of pta appears to be a futile attempt of cells to produce acetate as the major product. Instead, a cellular energy limitation resulted in the accumulation of intracellular pyruvate and upregulation of Pfl to convert formate to pyruvate. Therefore, metabolism is controlled, at least partially, at the protein expression level, an unusual feature for an acetogen. We anticipate that formate could be an important one-carbon substrate for acetogens to produce chemicals rich in pyruvate, a metabolite generally in low abundance during syngas growth.

Constructing a library of thousands of single-gene knockout or interference strains is a powerful tool to understand the relation between genotype and phenotype, but it is labor and cost intensive. Powered by the computer-aided gene annotation and functional grouping of non-essential genes, we showed that targeting a single gene directly to a specific observed phenotype could be quickly achieved for a specific microorganism via constructing a library of strains containing single chromosome-segment-deletion per strain. As a proof-of-concept, a genome-scale library consisting of 70 chromosome-segment-deletion strains for B. subtilis was constructed by CRISPR-based methods and strains with six loss-of- and gain-of-function phenotypes were screened out. To facilitate the rapid genotyping, we developed a web tool to visualize the potential targets of each chromosome segment associated with a particular function, successfully identifying the genes for valuable representative phenotypes. To apply the library to metabolic engineering, the hosts with improved production capacity of acetoin and lycopene were screened in the presence of pathway genes. This work demonstrated the significance of our strategy of chromosome segment scanning for gain- or loss-of-function screening (CHASING) on functional genomics investigation, robust chassis engineering, and chemical overproduction. O_FIG O_LINKSMALLFIG WIDTH=159 HEIGHT=200 SRC="FIGDIR/small/578163v1_ufig1.gif" ALT="Figure 1"> View larger version (36K): org.highwire.dtl.DTLVardef@19b4938org.highwire.dtl.DTLVardef@1e41d2eorg.highwire.dtl.DTLVardef@137d4b3org.highwire.dtl.DTLVardef@6d4bfc_HPS_FORMAT_FIGEXP M_FIG TOC C_FIG

D-Xylose, a major constituent of plant biomass and second most abundant sugar on Earth, holds a considerable potential as a substrate for sustainable bio-production. Pseudomonas putida KT2440 is an attractive bacterial host for valorizing biogenic feedstocks but lacks a xylose utilization pathway. While several attempts to engineer P. putida for growth on xylose have been reported, a comprehensive understanding of xylose metabolism in this bacterium is lacking, hindering its further improvement and rational tailoring for specific biotechnological purposes. In this study, we elucidated the xylose metabolism in the genome-reduced P. putida strain, EM42, endowed with xylose isomerase pathway (xylAB) and transporter (xylE) from Escherichia coli and used the obtained knowledge in combination with adaptive laboratory evolution to accelerate the bacteriums growth on the pentose sugar. Carbon flux analyses, targeted gene knock-outs, and in vitro enzyme assays portrayed xylose assimilation in P. putida and confirmed a partially cyclic upper xylose metabolism. Deletion of the local transcriptional regulator gene hexR de-repressed genes of several key catabolic enzymes and reduced the lag phase on xylose. Guided by metabolic modeling, we augmented P. putida with additional heterologous pentose phosphate pathway genes and subjected rationally prepared strains to adaptive laboratory evolution (ALE) on xylose. The descendants showed accelerated growth and reduced growth lag. Genomic and proteomic analysis of engineered and evolved mutants revealed the importance of a large genomic re-arrangement, transaldolase overexpression, and balancing gene expression in the synthetic xylABE operon. Importantly, omics analyses found that similar growth characteristics of two superior mutants were achieved through distinct evolutionary paths. This work provides a unique insight into how cell metabolism adjusts to a non-native substrate; it highlights the remarkable genomic and metabolic plasticity of P. putida and demonstrates the power of combining knowledge-driven engineering with ALE in generating desirable microbial phenotypes. HighlightsO_LIElucidated xylose catabolism via exogenous isomerase pathway in P. putida EM42. C_LIO_LIDeletion of transcriptional regulator HexR improved growth on xylose. C_LIO_LIKnowledge-guided interventions and adaptive evolution accelerated growth. C_LIO_LIOmics analyses of selected mutants highlighted the genomic and metabolic plasticity of P. putida. C_LIO_LITwo mutants with superior characteristics emerged from distinct evolutionary paths. C_LI