Engineering the carbon and redox metabolism of Paenibacillus polymyxa for efficient isobutanol production.

Paenibacillus polymyxa is a non-pathogenic, Gram-positive bacterium endowed with a rich and versatile metabolism. However interesting, this bacterium has been seldom used for bioproduction thus far. In this study, we engineered P. polymyxa for isobutanol production, a relevant bulk chemical and next-generation biofuel. A CRISPR-Cas9-based genome editing tool facilitated the chromosomal integration of a synthetic operon to establish isobutanol production. The 2,3-butanediol biosynthesis pathway, leading to the main fermentation product of P. polymyxa, was eliminated. A mutant strain harbouring the synthetic isobutanol operon (kdcA from Lactococcus lactis, and the native ilvC, ilvD and adh genes) produced 1 g L-1 isobutanol under microaerobic conditions. Improving NADPH regeneration by overexpression of the malic enzyme subsequently increased the product titre by 50%. Network-wide proteomics provided insights into responses of P. polymyxa to isobutanol and revealed a significant metabolic shift caused by alcohol production. Glucose-6-phosphate 1-dehydrogenase, the key enzyme in the pentose phosphate pathway, was identified as a bottleneck that hindered efficient NADPH regeneration through this pathway. Furthermore, we conducted culture optimization towards cultivating P. polymyxa in a synthetic minimal medium. We identified biotin (B7), pantothenate (B5) and folate (B9) to be mutual essential vitamins for P. polymyxa. Our rational metabolic engineering of P. polymyxa for the production of a heterologous chemical sheds light on the metabolism of this bacterium towards further biotechnological exploitation.

Time-resolved, deuterium-based fluxomics uncovers the hierarchy and dynamics of sugar processing by Pseudomonas putida.

Pseudomonas putida, a microbial host widely adopted for metabolic engineering, processes glucose through convergent peripheral pathways that ultimately yield 6-phosphogluconate. The periplasmic gluconate shunt (PGS), composed by glucose and gluconate dehydrogenases, sequentially transforms glucose into gluconate and 2-ketogluconate. Although the secretion of these organic acids by P. putida has been extensively recognized, the mechanism and spatiotemporal regulation of the PGS remained elusive thus far. To address this challenge, we adopted a dynamic 13C- and 2H-metabolic flux analysis strategy, termed D-fluxomics. D-fluxomics demonstrated that the PGS underscores a highly dynamic metabolic architecture in glucose-dependent batch cultures of P. putida, characterized by hierarchical carbon uptake by the PGS throughout the cultivation. Additionally, we show that gluconate and 2-ketogluconate accumulation and consumption can be solely explained as a result of the interplay between growth rate-coupled and decoupled metabolic fluxes. As a consequence, the formation of these acids in the PGS is inversely correlated to the bacterial growth rate-unlike the widely studied overflow metabolism of Escherichia coli and yeast. Our findings, which underline survival strategies of soil bacteria thriving in their natural environments, open new avenues for engineering P. putida towards efficient, sugar-based bioprocesses.

Emergent CRISPR-Cas-based technologies for engineering non-model bacteria.

Clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated proteins (Cas) technologies brought a transformative change in the way bacterial genomes are edited, and a plethora of studies contributed to developing multiple tools based on these approaches. Prokaryotic biotechnology benefited from the implementation of such genome engineering strategies, with an increasing number of non-model bacterial species becoming genetically tractable. In this review, we summarize the recent trends in engineering non-model microbes using CRISPR-Cas technologies, discussing their potential in supporting cell factory design towards biotechnological applications. These efforts include, among other examples, genome modifications as well as tunable transcriptional regulation (both positive and negative). Moreover, we examine how CRISPR-Cas toolkits for engineering non-model organisms enabled the exploitation of emergent biotechnological processes (e.g. native and synthetic assimilation of one-carbon substrates). Finally, we discuss our slant on the future of bacterial genome engineering for domesticating non-model organisms in light of the most recent advances in the ever-expanding CRISPR-Cas field.

Automating the design-build-test-learn cycle towards next-generation bacterial cell factories.

Automation is playing an increasingly significant role in synthetic biology. Groundbreaking technologies, developed over the past 20 years, have enormously accelerated the construction of efficient microbial cell factories. Integrating state-of-the-art tools (e.g. for genome engineering and analytical techniques) into the design-build-test-learn cycle (DBTLc) will shift the metabolic engineering paradigm from an almost artisanal labor towards a fully automated workflow. Here, we provide a perspective on how a fully automated DBTLc could be harnessed to construct the next-generation bacterial cell factories in a fast, high-throughput fashion. Innovative toolsets and approaches that pushed the boundaries in each segment of the cycle are reviewed to this end. We also present the most recent efforts on automation of the DBTLc, which heralds a fully autonomous pipeline for synthetic biology in the near future.

SEVA 4.0: an update of the Standard European Vector Architecture database for advanced analysis and programming of bacterial phenotypes.

The SEVA platform (https://seva-plasmids.com) was launched one decade ago, both as a database (DB) and as a physical repository of plasmid vectors for genetic analysis and engineering of Gram-negative bacteria with a structure and nomenclature that follows a strict, fixed architecture of functional DNA segments. While the current update keeps the basic features of earlier versions, the platform has been upgraded not only with many more ready-to-use plasmids but also with features that expand the range of target species, harmonize DNA assembly methods and enable new applications. In particular, SEVA 4.0 includes (i) a sub-collection of plasmids for easing the composition of multiple DNA segments with MoClo/Golden Gate technology, (ii) vectors for Gram-positive bacteria and yeast and [iii] off-the-shelf constructs with built-in functionalities. A growing collection of plasmids that capture part of the standard-but not its entirety-has been compiled also into the DB and repository as a separate corpus (SEVAsib) because of its value as a resource for constructing and deploying phenotypes of interest. Maintenance and curation of the DB were accompanied by dedicated diffusion and communication channels that make the SEVA platform a popular resource for genetic analyses, genome editing and bioengineering of a large number of microorganisms.

Engineering of Pseudomonas putida for accelerated co-utilization of glucose and cellobiose yields aerobic overproduction of pyruvate explained by an upgraded metabolic model.

Pseudomonas putida KT2440 is an attractive bacterial host for biotechnological production of valuable chemicals from renewable lignocellulosic feedstocks as it can valorize lignin-derived aromatics or glucose obtainable from cellulose. P. putida EM42, a genome-reduced variant of strain KT2440 endowed with advantageous physiological properties, was recently engineered for growth on cellobiose, a major cellooligosaccharide product of enzymatic cellulose hydrolysis. Co-utilization of cellobiose and glucose was achieved in a mutant lacking periplasmic glucose dehydrogenase Gcd (PP_1444). However, the cause of the co-utilization phenotype remained to be understood and the Δgcd strain had a significant growth defect. In this study, we investigated the basis of the simultaneous uptake of the two sugars and accelerated the growth of P. putida EM42 Δgcd mutant for the bioproduction of valuable compounds from glucose and cellobiose. We show that the gcd deletion lifted the inhibition of the exogenous ß-glucosidase BglC from Thermobifida fusca exerted by the intermediates of the periplasmic glucose oxidation pathway. The additional deletion of hexR gene, which encodes a repressor of the upper glycolysis genes, failed to restore rapid growth on glucose. The reduced growth rate of the Δgcd mutant was partially compensated by the implantation of heterologous glucose and cellobiose transporters (Glf from Zymomonas mobilis and LacY from Escherichia coli, respectively). Remarkably, this intervention resulted in the accumulation of pyruvate in aerobic P. putida cultures. We demonstrated that the excess of this key metabolic intermediate can be redirected to the enhanced biosynthesis of ethanol and lactate. The pyruvate overproduction phenotype was then unveiled by an upgraded genome-scale metabolic model constrained with proteomic and kinetic data. The model pointed to the saturation of glucose catabolism enzymes due to unregulated substrate uptake and it predicted improved bioproduction of pyruvate-derived chemicals by the engineered strain. This work sheds light on the co-metabolism of cellulosic sugars in an attractive biotechnological host and introduces a novel strategy for pyruvate overproduction in bacterial cultures under aerobic conditions.

A Nonconventional Archaeal Fluorinase Identified by In Silico Mining for Enhanced Fluorine Biocatalysis.

Fluorinases, the only enzymes known to catalyze the transfer of fluorine to an organic molecule, are essential catalysts for the biological synthesis of valuable organofluorines. However, the few fluorinases identified so far have low turnover rates that hamper biotechnological applications. Here, we isolated and characterized putative fluorinases retrieved from systematic in silico mining and identified a nonconventional archaeal enzyme from Methanosaeta sp. that mediates the fastest SN2 fluorination rate reported to date. Furthermore, we demonstrate enhanced production of fluoronucleotides in vivo in a bacterial host engineered with this archaeal fluorinase, paving the way toward synthetic metabolism for efficient biohalogenation.

Modular (de)construction of complex bacterial phenotypes by CRISPR/nCas9-assisted, multiplex cytidine base-editing.

CRISPR/Cas technologies constitute a powerful tool for genome engineering, yet their use in non-traditional bacteria depends on host factors or exogenous recombinases, which limits both efficiency and throughput. Here we mitigate these practical constraints by developing a widely-applicable genome engineering toolset for Gram-negative bacteria. The challenge is addressed by tailoring a CRISPR base editor that enables single-nucleotide resolution manipulations (C·G → T·A) with >90% efficiency. Furthermore, incorporating Cas6-mediated processing of guide RNAs in a streamlined protocol for plasmid assembly supports multiplex base editing with >85% efficiency. The toolset is adopted to construct and deconstruct complex phenotypes in the soil bacterium Pseudomonas putida. Single-step engineering of an aromatic-compound production phenotype and multi-step deconstruction of the intricate redox metabolism illustrate the versatility of multiplex base editing afforded by our toolbox. Hence, this approach overcomes typical limitations of previous technologies and empowers engineering programs in Gram-negative bacteria that were out of reach thus far.

Merging automation and fundamental discovery into the design-build-test-learn cycle of nontraditional microbes.

Major advances toward a bio-based industry enabled cost-efficient bioproduction of multiple chemicals, yet successful industrial processes are relatively scarce and limited to the use of few workhorse microbes as hosts. An in-depth understanding of the physiology and metabolism of nontraditional microorganisms is key to unleash their biotechnological potential. The inception of biofoundries multiplied the capacity of constructing and testing a large number of microbial strains tailored for bioproduction - and we argue that automation workflows therein can be adapted to gain fundamental knowledge of nontraditional hosts. Here, we propose a 'metabolism-centric' approach to the design-build-test-learn cycle of synthetic biology, supported by multi-omic analyses, to facilitate the deployment of microbial cell factories designed for bioproduction beyond the typical landscape of target products.

Model-guided dynamic control of essential metabolic nodes boosts acetyl-coenzyme A-dependent bioproduction in rewired Pseudomonas putida.

Pseudomonas putida is evolutionarily endowed with features relevant for bioproduction, especially under harsh operating conditions. The rich metabolic versatility of this species, however, comes at the price of limited formation of acetyl-coenzyme A (CoA) from sugar substrates. Since acetyl-CoA is a key metabolic precursor for a number of added-value products, in this work we deployed an in silico-guided rewiring program of central carbon metabolism for upgrading P. putida as a host for acetyl-CoA-dependent bioproduction. An updated kinetic model, integrating fluxomics and metabolomics datasets in addition to manually-curated information of enzyme mechanisms, identified targets that would lead to increased acetyl-CoA levels. Based on these predictions, a set of plasmids based on clustered regularly interspaced short palindromic repeats (CRISPR) and dead CRISPR-associated protein 9 (dCas9) was constructed to silence genes by CRISPR interference (CRISPRi). Dynamic reduction of gene expression of two key targets (gltA, encoding citrate synthase, and the essential accA gene, encoding subunit A of the acetyl-CoA carboxylase complex) mediated an 8-fold increase in the acetyl-CoA content of rewired P. putida. Poly(3-hydroxybutyrate) (PHB) was adopted as a proxy of acetyl-CoA availability, and two synthetic pathways were engineered for biopolymer accumulation. By including cell morphology as an extra target for the CRISPRi approach, fully rewired P. putida strains programmed for PHB accumulation had a 5-fold increase in PHB titers in bioreactor cultures using glucose. Thus, the strategy described herein allowed for rationally redirecting metabolic fluxes in P. putida from central metabolism towards product biosynthesis-especially relevant when deletion of essential pathways is not an option.

Pseudomonas taiwanensis biofilms for continuous conversion of cyclohexanone in drip flow and rotating bed reactors.

In this study, the biocatalytic performance of a Baeyer-Villiger monooxygenase (BVMO) catalyzing the reaction of cyclohexanone to ε-caprolactone was investigated in Pseudomonas biofilms. Biofilm growth and development of two Pseudomonas taiwanensis VLB120 variants, Ps_BVMO and Ps_BVMO_DGC, were evaluated in drip flow reactors (DFRs) and rotating bed reactors (RBRs). Engineering a hyperactive diguanylate cyclase (DGC) from Caulobacter crescentus into Ps_BVMO resulted in faster biofilm growth compared to the control Ps_BVMO strain in the DFRs. The maximum product formation rates of 92 and 87 g m-2 d-1 were observed for mature Ps_BVMO and Ps_ BVMO_DGC biofilms, respectively. The application of the engineered variants in the RBR was challenged by low biofilm surface coverage (50-60%) of rotating bed cassettes, side-products formation, oxygen limitation, and a severe drop in production rates with time. By implementing an active oxygen supply mode and a twin capillary spray feed, the biofilm surface coverage was maximized to 70-80%. BVMO activity was severely inhibited by cyclohexanol formation, resulting in a decrease in product formation rates. By controlling the cyclohexanone feed concentration at 4 mM, a stable product formation rate of 14 g m-2 d-1 and a substrate conversion of 60% was achieved in the RBR.

Rapid Genome Engineering of Pseudomonas Assisted by Fluorescent Markers and Tractable Curing of Plasmids.

Precise genome engineering has become a commonplace technique for metabolic engineering. Also, insertion, deletion and alteration of genes and other functional DNA sequences are essential for understanding and engineering cells. Several techniques have been developed to this end (e.g., CRISPR/Cas-assisted methods, homologous recombination, or λ Red recombineering), yet most of them rely on the use of auxiliary plasmids, which have to be cured after the editing procedure. Temperature-sensitive replicons, counter-selectable markers or repeated passaging of plasmid-bearing cells have been traditionally employed to circumvent this hurdle. While these protocols work reasonably well in some bacteria, they are not applicable for other species or are time consuming and laborious. Here, we present a fast and versatile protocol of fluorescent marker-assisted genome editing in Pseudomonas putida, followed by clean curing of auxiliary plasmids through user-controlled plasmid replication. One fluorescent marker facilitates identification of genome-edited colonies, while the second reporter enables detection of plasmid-free bacterial clones. Not only is this protocol the fastest available for Pseudomonas species, but it can be easily adapted to any type of genome modifications, including sequence deletions, insertions, and replacements. Graphical abstract: Rapid genome engineering of Pseudomonas with curable plasmids.

Negative regulation of plastidial isoprenoid pathway by herbivore-induced ß-cyclocitral in Arabidopsis thaliana.

Insect damage to plants is known to up-regulate defense and down-regulate growth processes. While there are frequent reports about up-regulation of defense signaling and production of defense metabolites in response to herbivory, much less is understood about the mechanisms by which growth and carbon assimilation are down-regulated. Here we demonstrate that insect herbivory down-regulates the 2-C-methyl-D-erythritol-4-phosphate (MEP) pathway in Arabidopsis (Arabidopsis thaliana), a pathway making primarily metabolites for use in photosynthesis. Simulated feeding by the generalist herbivore Spodoptera littoralis suppressed flux through the MEP pathway and decreased steady-state levels of the intermediate 1-deoxy-D-xylulose 5-phosphate (DXP). Simulated herbivory also increased reactive oxygen species content which caused the conversion of ß-carotene to ß-cyclocitral (ßCC). This volatile oxidation product affected the MEP pathway by directly inhibiting DXP synthase (DXS), the rate-controlling enzyme of the MEP pathway in Arabidopsis and inducing plant resistance against S. littoralis ßCC inhibited both DXS transcript accumulation and DXS activity. Molecular models suggested that ßCC binds to DXS at the binding site for the thymine pyrophosphate cofactor and blocks catalysis, which was confirmed by direct assays of ßCC with the purified DXS protein in vitro. Another intermediate of the MEP pathway, 2-C-methyl-D-erythritol-2, 4-cyclodiphosphate, which is known to stimulate salicylate defense signaling, showed greater accumulation and enhanced export out of the plastid in response to simulated herbivory. Together, our work implicates ßCC as a signal of herbivore damage in Arabidopsis that increases defense and decreases flux through the MEP pathway, a pathway involved in growth and carbon assimilation.

A fluoride-responsive genetic circuit enables in vivo biofluorination in engineered Pseudomonas putida.

Fluorine is a key element in the synthesis of molecules broadly used in medicine, agriculture and materials. Addition of fluorine to organic structures represents a unique strategy for tuning molecular properties, yet this atom is rarely found in Nature and approaches to integrate fluorometabolites into the biochemistry of living cells are scarce. In this work, synthetic gene circuits for organofluorine biosynthesis are implemented in the platform bacterium Pseudomonas putida. By harnessing fluoride-responsive riboswitches and the orthogonal T7 RNA polymerase, biochemical reactions needed for in vivo biofluorination are wired to the presence of fluoride (i.e. circumventing the need of feeding expensive additives). Biosynthesis of fluoronucleotides and fluorosugars in engineered P. putida is demonstrated with mineral fluoride both as only fluorine source (i.e. substrate of the pathway) and as inducer of the synthetic circuit. This approach expands the chemical landscape of cell factories by providing alternative biosynthetic strategies towards fluorinated building-blocks.

Industrial biotechnology of Pseudomonas putida: advances and prospects.

Pseudomonas putida is a Gram-negative, rod-shaped bacterium that can be encountered in diverse ecological habitats. This ubiquity is traced to its remarkably versatile metabolism, adapted to withstand physicochemical stress, and the capacity to thrive in harsh environments. Owing to these characteristics, there is a growing interest in this microbe for industrial use, and the corresponding research has made rapid progress in recent years. Hereby, strong drivers are the exploitation of cheap renewable feedstocks and waste streams to produce value-added chemicals and the steady progress in genetic strain engineering and systems biology understanding of this bacterium. Here, we summarize the recent advances and prospects in genetic engineering, systems and synthetic biology, and applications of P. putida as a cell factory. KEY POINTS: • Pseudomonas putida advances to a global industrial cell factory. • Novel tools enable system-wide understanding and streamlined genomic engineering. • Applications of P. putida range from bioeconomy chemicals to biosynthetic drugs.

Synthetic control of plasmid replication enables target- and self-curing of vectors and expedites genome engineering of Pseudomonas putida.

Genome engineering of non-conventional microorganisms calls for the development of dedicated synthetic biology tools. Pseudomonas putida is a Gram-negative, non-pathogenic soil bacterium widely used for metabolic engineering owing to its versatile metabolism and high levels of tolerance to different types of stress. Genome editing of P. putida largely relies on homologous recombination events, assisted by helper plasmid-based expression of genes encoding DNA modifying enzymes. Plasmid curing from selected isolates is the most tedious and time-consuming step of this procedure, and implementing commonly used methods to this end in P. putida (e.g. temperature-sensitive replicons) is often impractical. To tackle this issue, we have developed a toolbox for both target- and self-curing of plasmid DNA in Pseudomonas species. Our method enables plasmid-curing in a simple cultivation step by combining in vivo digestion of vectors by the I-SceI homing nuclease with synthetic control of plasmid replication, triggered by the addition of a cheap chemical inducer (3-methylbenzoate) to the medium. The system displays an efficiency of vector curing >90% and the screening of plasmid-free clones is greatly facilitated by the use of fluorescent markers that can be selected according to the application intended. Furthermore, quick genome engineering of P. putida using self-curing plasmids is demonstrated through genome reduction of the platform strain EM42 by eliminating all genes encoding ß-lactamases, the catabolic ben gene cluster, and the pyoverdine synthesis machinery. Physiological characterization of the resulting streamlined strain, P. putida SEM10, revealed advantageous features that could be exploited for metabolic engineering.

An expanded CRISPRi toolbox for tunable control of gene expression in Pseudomonas putida.

Owing to its wide metabolic versatility and physiological robustness, together with amenability to genetic manipulations and high resistance to stressful conditions, Pseudomonas putida is increasingly becoming the organism of choice for a range of applications in both industrial and environmental applications. However, a range of applied synthetic biology and metabolic engineering approaches are still limited by the lack of specific genetic tools to effectively and efficiently regulate the expression of target genes. Here, we present a single-plasmid CRISPR-interference (CRISPRi) system expressing a nuclease-deficient cas9 gene under the control of the inducible XylS/Pm expression system, along with the option of adopting constitutively expressed guide RNAs (either sgRNA or crRNA and tracrRNA). We showed that the system enables tunable, tightly controlled gene repression (up to 90%) of chromosomally expressed genes encoding fluorescent proteins, either individually or simultaneously. In addition, we demonstrate that this method allows for suppressing the expression of the essential genes pyrF and ftsZ, resulting in significantly low growth rates or morphological changes respectively. This versatile system expands the capabilities of the current CRISPRi toolbox for efficient, targeted and controllable manipulation of gene expression in P. putida.

Physical decoupling of XylS/Pm regulatory elements and conditional proteolysis enable precise control of gene expression in Pseudomonas putida.

Most of the gene expression systems available for Gram-negative bacteria are afflicted by relatively high levels of basal (i.e. leaky) expression of the target gene(s). This occurrence affects the system dynamics, ultimately reducing the output and productivity of engineered pathways and synthetic circuits. In order to circumvent this problem, we have designed a novel expression system based on the well-known XylS/Pm transcriptional regulator/promoter pair from the soil bacterium Pseudomonas putida mt-2, in which the key functional elements are physically decoupled. By integrating the xylS gene into the chromosome of the platform strain KT2440, while placing the Pm promoter into a set of standard plasmid vectors, the inducibility of the system (i.e. the output difference between the induced and uninduced state) improved up to 170-fold. We further combined this modular system with an extra layer of post-translational control by means of conditional proteolysis. In this setup, the target gene is tagged with a synthetic motif dictating protein degradation. When the system features were characterized using the monomeric superfolder GFP as a model protein, the basal levels of fluorescence were brought down to zero (i.e. below the limit of detection). In all, these novel expression systems constitute an alternative tool to altogether suppress leaky gene expression, and they can be easily adapted to other vector formats and plugged-in into different Gram-negative bacterial species at the user's will.

Non-invasive, ratiometric determination of intracellular pH in Pseudomonas species using a novel genetically encoded indicator.

The ability of Pseudomonas species to thrive in all major natural environments (i.e. terrestrial, freshwater and marine) is based on its exceptional capability to adapt to physicochemical changes. Thus, environmental bacteria have to tightly control the maintenance of numerous physiological traits across different conditions. The intracellular pH (pHi ) homoeostasis is a particularly important feature, since the pHi influences a large portion of the biochemical processes in the cell. Despite its importance, relatively few reliable, easy-to-implement tools have been designed for quantifying in vivo pHi changes in Gram-negative bacteria with minimal manipulations. Here we describe a convenient, non-invasive protocol for the quantification of the pHi in bacteria, which is based on the ratiometric fluorescent indicator protein PHP (pH indicator for Pseudomonas). The DNA sequence encoding PHP was thoroughly adapted to guarantee optimal transcription and translation of the indicator in Pseudomonas species. Our PHP-based quantification method demonstrated that pHi is tightly regulated over a narrow range of pH values not only in Pseudomonas, but also in other Gram-negative bacterial species such as Escherichia coli. The maintenance of the cytoplasmic pH homoeostasis in vivo could also be observed upon internal (e.g. redirection of glucose consumption pathways in P. putida) and external (e.g. antibiotic exposure in P. aeruginosa) perturbations, and the PHP indicator was also used to follow dynamic changes in the pHi upon external pH shifts. In summary, our work describes a reliable method for measuring pHi in Pseudomonas, allowing for the detailed investigation of bacterial pHi homoeostasis and its regulation.