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

Comparing three emerging industrial cell factories: Pseudomonas putida KT2440, Halomonas bluephagenesis TD01, and Zymomonas mobilis ZM4

Nonmodel microbes with unique advantages are emerging as industrial platforms, driven by advances in genetic engineering and omics technologies. Notable examples include the versatile soil bacterium Pseudomonas putida KT2440, the halophilic Halomonas bluephagenesis TD01, and the ethanologenic Zymomonas mobilis ZM4. While all three primarily use the Entner-Doudoroff pathway for glucose metabolism, they differ in various metabolic pathways and product synthesis. This review summarizes and compares their central carbon metabolism, advancements in genome engineering tools, and progress in scaling industrial applications from lab scale, to pilot scale, to full-scale commercial production. Understanding their similarities and differences informs future research on optimizing industrial applications and may guide the development of new microbial hosts.

Genetic Dissection of Cyclic di-GMP Signalling in Pseudomonas aeruginosa via Systematic Diguanylate Cyclase Disruption

The second messenger bis-(3' → 5')-cyclic dimeric guanosine monophosphate (c-di-GMP) governs adaptive responses in the opportunistic pathogen Pseudomonas aeruginosa, including biofilm formation and the transition from acute to chronic infections. Understanding the intricate c-di-GMP signalling network remains challenging due to the overlapping activities of numerous diguanylate cyclases (DGCs). In this study, we employed a CRISPR-based multiplex genome-editing tool to disrupt all 32 GGDEF domain-containing proteins (GCPs) implicated in c-di-GMP signalling in P. aeruginosa PA14. Phenotypic and physiological analyses revealed that the resulting mutant was unable to form biofilms and had attenuated virulence. Residual c-di-GMP levels were still detected despite the extensive GCP disruption, underscoring the robustness of this regulatory network. Taken together, these findings provide insights into the complex c-di-GMP metabolism and showcase the importance of functional overlapping in bacterial signalling. Moreover, our approach overcomes the native redundancy in c-di-GMP synthesis, providing a framework to dissect individual DGC functions and paving the way for targeted strategies to address bacterial adaptation and pathogenesis.

Computation-aided designs enable developing auxotrophic metabolic sensors for wide-range glyoxylate and glycolate detection

Auxotrophic metabolic sensors (AMS) are microbial strains modified so that biomass formation correlates with the availability of specific metabolites. These sensors are essential for bioengineering (e.g., in growth-coupled designs) but creating them is often a time-consuming and low-throughput process that can be streamlined by in silico analysis. Here, we present a systematic workflow for designing, implementing, and testing versatile AMS based on Escherichia coli. Glyoxylate, a key metabolite in (synthetic) CO2 fixation and carbon-conserving pathways, served as the test analyte. Through iterative screening of a compact metabolic model, we identify non-trivial growth-coupled designs that result in six AMS with a wide sensitivity range for glyoxylate, spanning three orders of magnitude in the detected analyte concentration. We further adapt these E. coli AMS for sensing glycolate and demonstrate their utility in both pathway engineering (testing a key metabolic module for carbon assimilation via glyoxylate) and environmental monitoring (quantifying glycolate produced by photosynthetic microalgae). Adapting this workflow to the sensing of different metabolites could facilitate the design and implementation of AMS for diverse biotechnological applications.

CIFR (Clone-Integrate-Flip-out-Repeat): A toolset for iterative genome and pathway engineering of Gram-negative bacteria

Advanced genome engineering enables precise and customizable modifications of bacterial species, and toolsets that exhibit broad-host compatibility are particularly valued owing to their portability. Tn5 transposon vectors have been widely used to establish random integrations of desired DNA sequences into bacterial genomes. However, the iteration of the procedure remains challenging because of the limited availability and reusability of selection markers. We addressed this challenge with CIFR, a mini-Tn5 integration system tailored for iterative genome engineering. The pCIFR vectors incorporate attP and attB sites flanking an antibiotic resistance marker used to select for the insertion. Subsequent removal of antibiotic determinants is facilitated by the Bxb1 integrase paired to a user-friendly counter-selection marker, both encoded in auxiliary plasmids. CIFR delivers engineered strains harboring stable DNA insertions and free of any antibiotic resistance cassette, allowing for the reusability of the tool. The system was validated in Pseudomonas putida, Escherichia coli, and Cupriavidus necator, underscoring its portability across diverse industrially relevant hosts. The CIFR toolbox was calibrated through combinatorial integrations of chromoprotein genes in P. putida, generating strains displaying a diverse color palette. We also introduced a carotenoid biosynthesis pathway in P. putida in a two-step engineering process, showcasing the potential of the tool for pathway balancing. The broad utility of the CIFR toolbox expands the toolkit for metabolic engineering, allowing for the construction of complex phenotypes while opening new possibilities in bacterial genetic manipulations.

Upcycling of polyamides through chemical hydrolysis and engineered Pseudomonas putida

Aliphatic polyamides, or nylons, are widely used in the textile and automotive industry due to their high durability and tensile strength, but recycling rates are below 5%. Chemical recycling of polyamides is possible but typically yields mixtures of monomers and oligomers which hinders downstream purification. Here, Pseudomonas putida KT2440 was engineered to metabolize C6-polyamide monomers such as 6-aminohexanoic acid, ε-caprolactam and 1,6-hexamethylenediamine, guided by adaptive laboratory evolution. Heterologous expression of nylonases also enabled P. putida to metabolize linear and cyclic nylon oligomers derived from chemical polyamide hydrolysis. RNA sequencing and reverse engineering revealed the metabolic pathways for these non-natural substrates. To demonstrate microbial upcycling, the phaCAB operon from Cupriavidus necator was heterologously expressed to enable production of polyhydroxybutyrate (PHB) from PA6 hydrolysates. This study presents a microbial host for the biological conversion, in combination with chemical hydrolysis, of polyamide monomers and mixed polyamids hydrolysates to a value-added product.

Precision engineering of the probiotic Escherichia coli Nissle 1917 with prime editing

CRISPR-Cas systems are transforming precision medicine with engineered probiotics as next-generation diagnostics and therapeutics. To promote human health and treat disease, engineering probiotic bacteria demands maximal versatility to enable non-natural functionalities while minimizing undesired genomic interferences. Here, we present a streamlined prime editing approach tailored for probiotic Escherichia coli Nissle 1917 utilizing only essential genetic modules, including Cas9 nickase from Streptococcus pyogenes, a codon-optimized reverse transcriptase, and a prime editing guide RNA, and an optimized workflow with longer induction. As a result, we achieved all types of prime editing in every individual round of experiments with efficiencies of 25.0%, 52.0%, and 66.7% for DNA deletion, insertion, and substitution, respectively. A comprehensive evaluation of off-target effects revealed a significant reduction in unintended mutations, particularly in comparison to two different base editing methods. Leveraging the prime editing system, we inserted a unique DNA sequence to barcode the edited strain and established an antibiotic-resistance-gene-free platform to enable non-natural functionalities. Our prime editing strategy presents a CRISPR-Cas system that can be readily implemented in any laboratories with the basic CRISPR setups, paving the way for future innovations in engineered probiotics.IMPORTANCEOne ultimate goal of gene editing is to introduce designed DNA variations at specific loci in living organisms with minimal unintended interferences in the genome. Achieving this goal is especially critical for creating engineered probiotics as living diagnostics and therapeutics to promote human health and treat diseases. In this endeavor, we report a customized prime editing system for precision engineering of probiotic Escherichia coli Nissle 1917. With such a system, we developed a barcoding system for tracking engineered strains, and we built an antibiotic-resistance-gene-free platform to enable non-natural functionalities. We provide not only a powerful gene editing approach for probiotic bacteria but also new insights into the advancement of innovative CRISPR-Cas systems.

Engineering a phi15-based expression system for stringent gene expression in Pseudomonas putida

The T7 phage RNA polymerase (RNAP) is a widely used expression platform, but its implementation in non-model microbial hosts poses significant challenges due to cytotoxicity. We constructed an optimized phage phi15-based expression system as alternative to the T7 platform for a wide range of applications in Pseudomonas putida. The new system employs the small phi15 RNAP, driving expression from an orthogonal phi15 promoter. By finetuning expression levels of phi15rnap and introducing a phi15 lysozyme mutant that inhibits phi15 RNAP in uninduced conditions, a stringent system was created with 200-fold inducibility. Moreover, by successfully decoupling cell growth and protein production using phi15 gp16, a host RNAP inhibitor, expression levels could be enhanced further (20%). Apart from creating four optimized platform P. putida hosts and a set of Golden Gate-compatible vectors, we demonstrate the extensive flexibility of the phi15 system. A proof-of-concept expression for industrially relevant fluorinase resulted in 2.5- and 5-fold increased yield compared to other widely-adopted expression systems. The system functions well in combination with several inducer systems, and in a variety of vector-based and genomically integrated set-ups. In conclusion, the phi15 RNAP, promoter, lysozyme and growth-decoupler provide a valuable plug-and-play set of genetic parts for the P. putida toolbox.

A universal and wide-range cytosine base editor via domain-inlaid and fidelity-optimized CRISPR-FrCas9

CRISPR-based base editor (BE) offer diverse editing options for genetic engineering of microorganisms, but its application is limited by protospacer adjacent motif (PAM) sequences, context preference, editing window, and off-target effects. Here, a series of iteratively improved cytosine base editors (CBEs) are constructed using the FrCas9 nickase (FrCas9n) with the unique PAM palindromic structure (NNTA) to alleviate these challenges. The deaminase domain-inlaid FrCas9n exhibits an editing range covering 38 nucleotides upstream and downstream of the palindromic PAM, without context preference, which is 6.3 times larger than that of traditional CBEs. Additionally, lower off-target editing is achieved when incorporating high-fidelity mutations at R61A and Q964A in FrCas9n, while maintaining high editing efficiency. The final CBE, HF-ID824-evoCDA-FrCas9n demonstrates broad applicability across different microbes such as Escherichia coli MG1655, Shewanella oneidensis MR-1, and Pseudomonas aeruginosa PAO1. Collectively, this tool offers robust gene editing for facilitating mechanistic studies, functional exploration, and protein evolution in microbes.

Leveraging Engineered Pseudomonas putida Minicells for Bioconversion of Organic Acids into Short-Chain Methyl Ketones

Methyl ketones, key building blocks widely used in diverse industrial applications, largely depend on oil-derived chemical methods for their production. Here, we investigated biobased production alternatives for short-chain ketones, adapting the solvent-tolerant soil bacterium Pseudomonas putida as a host for ketone biosynthesis either by whole-cell biocatalysis or using engineered minicells, chromosome-free bacterial vesicles. Organic acids (acetate, propanoate and butanoate) were selected as the main carbon substrate to drive the biosynthesis of acetone, butanone and 2-pentanone. Pathway optimization identified efficient enzyme variants from Clostridium acetobutylicum and Escherichia coli, tested with both constitutive and inducible expression of the cognate genes. By implementing these optimized pathways in P. putida minicells, which can be prepared through a simple three-step purification protocol, the feedstock was converted into the target short-chain methyl ketones. These results highlight the value of combining morphology and pathway engineering of noncanonical bacterial hosts to establish alternative bioprocesses for toxic chemicals that are difficult to produce by conventional approaches.

Accelerated Metabolic Engineering for Industrial Strain Development via the Construction of a Large-Scale Genome Library

Production of chemicals via metabolic engineering of microbes is becoming highly important for sustainable bioeconomy. Conventional metabolic engineering methodologies typically involve labor-intensive and time-consuming processes of iterative genetic modifications, which are inefficient in identifying new genetic targets for the construction of robust industrial strains on a large scale. To accelerate the creation of efficient microbial cell factories and enhance our insights into cellular metabolism, diverse large-scale genome libraries are emerging as powerful tools, which can be established through multiplex or parallel genome editing, gene expression regulation, and incorporation of evolutionary strategies. In this review, we discuss the latest advancements in the construction of genome-scale libraries as well as their applications within the domain of metabolic engineering. We also address the limitations of various techniques and provide insights into future prospects for the field.

Hypermutability bypasses genetic constraints in SCV phenotypic switching in Pseudomonas aeruginosa biofilms

Biofilms are critical in the persistence of Pseudomonas aeruginosa infections, particularly in cystic fibrosis patients. This study explores the adaptive mechanisms behind the phenotypic switching between Small Colony Variants (SCVs) and revertant states in P. aeruginosa biofilms, emphasizing hypermutability due to Mismatch Repair System (MRS) deficiencies. Through experimental evolution and whole-genome sequencing, we show that both wild-type and mutator strains undergo parallel evolution by accumulating compensatory mutations in factors regulating intracellular c-di-GMP levels, particularly in the Wsp and Yfi systems. While wild-type strains face genetic constraints, mutator strains bypass these by accessing alternative genetic pathways regulating c-di-GMP and biofilm formation. This increased genetic accessibility, driven by higher mutation rates and specific mutational biases, supports sustained cycles of SCV conversion and reversion. Our findings underscore the crucial role of hypermutability in P. aeruginosa adaptation, with significant implications for managing persistent infections in clinical settings.

Selection of Fusion-Site Overhang Sets for High-Fidelity and High-Complexity Golden Gate Assembly

Golden Gate Assembly depends on the accurate ligation of overhangs at fragment fusion sites to generate full-length products with all parts in the desired order. Traditionally, fusion-site sequences are selected by using validated sets of overhang sequences or by applying a handful of semi-empirical rules to guide overhang choice. While these approaches allow dependable assembly of 6-8 fragments in one pot, recent work has demonstrated that comprehensive measurement of ligase fidelity allows prediction of high-fidelity junction sets that enable much more complex assemblies of 12, 24, or even 36+ fragments in a single reaction that will join with high accuracy and efficiency. In this chapter, we outline the application of a set of online tools that apply these comprehensive datasets to the analysis of existing junction sets, the de novo selection of new high-fidelity overhang sets, the modification and expansion of existing sets, and the principles for dividing known sequences at an arbitrary number of high-fidelity breakpoints.

One-for-all gene inactivation via PAM-independent base editing in bacteria

Base editing is preferable for bacterial gene inactivation without generating double-strand breaks, requiring homology recombination, or highly efficient DNA delivery capability. However, the potential of base editing is limited by the adjoined dependence on the editing window and protospacer adjacent motif. Herein, we report an unconstrained base-editing system to enable the inactivation of any genes of interest in bacteria. We employed a dCas9 derivative, dSpRY, and activation-induced cytidine deaminase to build a protospacer adjacent motif-independent base editor. Then, we programmed the base editor to exclude the START codon of a gene of interest instead of introducing premature STOP codons to obtain a universal approach for gene inactivation, namely XSTART, with an overall efficiency approaching 100%. By using XSTART, we successfully manipulated the amino acid metabolisms in Escherichia coli, generating glutamine, arginine, and aspartate auxotrophic strains. While we observed a high frequency of off-target events as a trade-off for increased efficiency, refining the regulatory system of XSTART to limit expression levels reduced off-target events by over 60% without sacrificing efficiency, aligning our results with previously reported levels. Finally, the effectiveness of XSTART was also demonstrated in probiotic E. coli Nissle 1917 and photoautotrophic cyanobacterium Synechococcus elongatus, illustrating its potential in reprogramming diverse bacteria.

CRISPR-Cas9-based approaches for genetic analysis and epistatic interaction studies in Coxiella burnetii

Coxiella burnetii is an obligate intracellular bacterial pathogen that replicates to high numbers in an acidified lysosome-derived vacuole. Intracellular replication requires the Dot/Icm type IVB secretion system, which translocates over 100 different effector proteins into the host cell. Screens employing random transposon mutagenesis have identified several C. burnetii effectors that play an important role in intracellular replication; however, the difficulty in conducting directed mutagenesis has been a barrier to the systematic analysis of effector mutants and to the construction of double mutants to assess epistatic interactions between effectors. Here, two CRISPR-Cas9 technology-based approaches were developed to study C. burnetii phenotypes resulting from targeted gene disruptions. CRISPRi was used to silence gene expression and demonstrated that silencing of effectors or Dot/Icm system components resulted in phenotypes similar to those of transposon insertion mutants. A CRISPR-Cas9-mediated cytosine base editing protocol was developed to generate targeted loss-of-function mutants through the introduction of premature stop codons into C. burnetii genes. Cytosine base editing successfully generated double mutants in a single step. A double mutant deficient in both cig57 and cig2 had a robust and additive intracellular replication defect when compared to either single mutant, which is consistent with Cig57 and Cig2 functioning in independent pathways that both contribute to a vacuole that supports C. burnetii replication. Thus, CRISPR-Cas9-based technologies expand the genetic toolbox for C. burnetii and will facilitate genetic studies aimed at investigating the mechanisms this pathogen uses to replicate inside host cells.

IMPORTANCE

Understanding the genetic mechanisms that enable C. burnetii to replicate in mammalian host cells has been hampered by the difficulty in making directed mutations. Here, a reliable and efficient system for generating targeted loss-of-function mutations in C. burnetii using a CRISPR-Cas9-assisted base editing approach is described. This technology was applied to make double mutants in C. burnetii that enabled the genetic analysis of two genes that play independent roles in promoting the formation of vacuoles that support intracellular replication. This advance will accelerate the discovery of mechanisms important for C. burnetii host infection and disease.

Harnessing noncanonical redox cofactors to advance synthetic assimilation of one-carbon feedstocks

One-carbon (C1) feedstocks, such as carbon monoxide (CO), formate (HCO2H), methanol (CH3OH), and methane (CH4), can be obtained either through stepwise electrochemical reduction of CO2 with renewable electricity or via processing of organic side streams. These C1 substrates are increasingly investigated in biotechnology as they can contribute to a circular carbon economy. In recent years, noncanonical redox cofactors (NCRCs) emerged as a tool to generate synthetic electron circuits in cell factories to maximize electron transfer within a pathway of interest. Here, we argue that expanding the use of NCRCs in the context of C1-driven bioprocesses will boost product yields and facilitate challenging redox transactions that are typically out of the scope of natural cofactors due to inherent thermodynamic constraints.

The structural complexity of pyomelanin impacts UV shielding in Pseudomonas species with different lifestyles

Pyomelanin, a polymeric pigment in Pseudomonas, arises mainly from alterations in tyrosine degradation. The chemical structure of pyomelanin remains elusive due to its heterogeneous nature. Here, we report strain-specific differences in pyomelanin structural features across Pseudomonas using PAO1 and PA14 reference strains carrying mutations in hmgA (a gene involved in pyomelanin synthesis), a melanogenic P. aeruginosa clinical isolate (PAM), and a melanogenic P. extremaustralis (PexM). UV spectra showed dual peaks for PAO1 and PA14 mutants and single peaks for PAM and PexM. FTIR phenol : alcohol ratio changes and complex NMR spectra indicated non-linear polymers. UVC radiation survival increased with pyomelanin addition, correlating with pigment absorption attenuation. P. extremaustralis UVC survival varied with melanin source, with PAO1 pyomelanin being the most protective. These findings delineate structure-based pyomelanin subgroups, having distinct physiological effects.

Simultaneous multi-site editing of individual genomes using retron arrays

During recent years, the use of libraries-scale genomic manipulations scaffolded on CRISPR guide RNAs have been transformative. However, these existing approaches are typically multiplexed across genomes. Unfortunately, building cells with multiple, nonadjacent precise mutations remains a laborious cycle of editing, isolating an edited cell and editing again. The use of bacterial retrons can overcome this limitation. Retrons are genetic systems composed of a reverse transcriptase and a noncoding RNA that contains an multicopy single-stranded DNA, which is reverse transcribed to produce multiple copies of single-stranded DNA. Here we describe a technology-termed a multitron-for precisely modifying multiple sites on a single genome simultaneously using retron arrays, in which multiple donor-encoding DNAs are produced from a single transcript. The multitron architecture is compatible with both recombineering in prokaryotic cells and CRISPR editing in eukaryotic cells. We demonstrate applications for this approach in molecular recording, genetic element minimization and metabolic engineering.

Disentangling the Regulatory Response of Agrobacterium tumefaciens CHLDO to Glyphosate for Engineering Whole-Cell Phosphonate Biosensors

Phosphonates (PHTs), organic compounds with a stable C-P bond, are widely distributed in nature. Glyphosate (GP), a synthetic PHT, is extensively used in agriculture and has been linked to various human health issues and environmental damage. Given the prevalence of GP, developing cost-effective, on-site methods for GP detection is key for assessing pollution and reducing exposure risks. We adopted Agrobacterium tumefaciens CHLDO, a natural GP degrader, as a host and the source of genetic parts for constructing PHT biosensors. In this bacterial species, the phn gene cluster, encoding the C-P lyase pathway, is regulated by the PhnF transcriptional repressor. We selected the phnG promoter, which displays a dose-dependent response to GP, to build a set of whole-cell biosensors. Through stepwise genetic optimization of the transcriptional cascade, we created a whole-cell biosensor capable of detecting GP in the 0.25-50 μM range in various samples, including soil and water.

Accelerating enzyme discovery and engineering with high-throughput screening

Covering: up to August 2024Enzymes play an essential role in synthesizing value-added chemicals with high specificity and selectivity. Since enzymes utilize substrates derived from renewable resources, biocatalysis offers a pathway to an efficient bioeconomy with reduced environmental footprint. However, enzymes have evolved over millions of years to meet the needs of their host organisms, which often do not align with industrial requirements. As a result, enzymes frequently need to be tailored for specific industrial applications. Combining enzyme engineering with high-throughput screening has emerged as a key approach for developing novel biocatalysts, but several challenges are yet to be addressed. In this review, we explore emergent strategies and methods for isolating, creating, and characterizing enzymes optimized for bioproduction. We discuss fundamental approaches to discovering and generating enzyme variants and identifying those best suited for specific applications. Additionally, we cover techniques for creating libraries using automated systems and highlight innovative high-throughput screening methods that have been successfully employed to develop novel biocatalysts for natural product synthesis.

A versatile microbial platform as a tunable whole-cell chemical sensor

Biosensors are used to detect and quantify chemicals produced in industrial microbiology with high specificity, sensitivity, and portability. Most biosensors, however, are limited by the need for transcription factors engineered to recognize specific molecules. In this study, we overcome the limitations typically associated with traditional biosensors by engineering Pseudomonas putida for whole-cell sensing of a variety of chemicals. Our approach integrates fluorescent reporters with synthetic auxotrophies within central metabolism that can be complemented by target analytes in growth-coupled setups. This platform enables the detection of a wide array of structurally diverse chemicals under various conditions, including co-cultures of producer cell factories and sensor strains. We also demonstrate the applicability of this versatile biosensor platform for monitoring complex biochemical processes, including plastic degradation by either purified hydrolytic enzymes or engineered bacteria. This microbial system provides a rapid, sensitive, and readily adaptable tool for monitoring cell factory performance and for environmental analyzes.

CRISPR Tools for Engineering Prokaryotic Systems: Recent Advances and New Applications

In the past decades, the broad selection of CRISPR-Cas systems has revolutionized biotechnology by enabling multimodal genetic manipulation in diverse organisms. Rooted in a molecular engineering perspective, we recapitulate the different CRISPR components and how they can be designed for specific genetic engineering applications. We first introduce the repertoire of Cas proteins and tethered effectors used to program new biological functions through gene editing and gene regulation. We review current guide RNA (gRNA) design strategies and computational tools and how CRISPR-based genetic circuits can be constructed through regulated gRNA expression. Then, we present recent advances in CRISPR-based biosensing, bioproduction, and biotherapeutics across in vitro and in vivo prokaryotic systems. Finally, we discuss forthcoming applications in prokaryotic CRISPR technology that will transform synthetic biology principles in the near future.

Establishing a straightforward I-SceI-mediated recombination one-plasmid system for efficient genome editing in P. putida KT2440

Pseudomonas putida has become an increasingly important chassis for producing valuable bioproducts. This development is not least due to the ever-improving genetic toolbox, including gene and genome editing techniques. Here, we present a novel, one-plasmid design of a critical genetic tool, the pEMG/pSW system, guaranteeing one engineering cycle to be finalized in 3 days. The pEMG/pSW system proved in the last decade to be valuable for targeted genome engineering in Pseudomonas, as it enables the deletion of large regions of the genome, the integration of heterologous gene clusters or the targeted generation of point mutations. Here, to expedite genetic engineering, two alternative plasmids were constructed: (1) The sacB gene from Bacillus subtilis was integrated into the I-SceI expressing plasmid pSW-2 as a counterselection marker to accelerated plasmid curing; (2) double-strand break introducing gene I-sceI and sacB counterselection marker were integrated into the backbone of the original pEMG vector, named pEMG-RIS. The single plasmid of pEMG-RIS allows rapid genome editing despite the low transcriptional activity of a single copy of the I-SceI encoding gene. Here, the usability of the pEMG-RIS is shown in P. putida KT2440 by integrating an expression cassette including an msfGFP gene in 3 days. In addition, a large fragment of 12.1 kb was also integrated. In summary, we present an updated pEMG/pSW genome editing system that allows efficient and rapid genome editing in P. putida. All plasmids designed in this study will be available via the Addgene platform.

Multiplex CRISPR-Cas Genome Editing: Next-Generation Microbial Strain Engineering

Genome editing is a crucial technology for obtaining desired phenotypes in a variety of species, ranging from microbes to plants, animals, and humans. With the advent of CRISPR-Cas technology, it has become possible to edit the intended sequence by modifying the target recognition sequence in guide RNA (gRNA). By expressing multiple gRNAs simultaneously, it is possible to edit multiple targets at the same time, allowing for the simultaneous introduction of various functions into the cell. This can significantly reduce the time and cost of obtaining engineered microbial strains for specific traits. In this review, we investigate the resolution of multiplex genome editing and its application in engineering microorganisms, including bacteria and yeast. Furthermore, we examine how recent advancements in artificial intelligence technology could assist in microbial genome editing and engineering. Based on these insights, we present our perspectives on the future evolution and potential impact of multiplex genome editing technologies in the agriculture and food industry.

Enzymatic synthesis of mono- and trifluorinated alanine enantiomers expands the scope of fluorine biocatalysis

Fluorinated amino acids serve as an entry point for establishing new-to-Nature chemistries in biological systems, and novel methods are needed for the selective synthesis of these building blocks. In this study, we focused on the enzymatic synthesis of fluorinated alanine enantiomers to expand fluorine biocatalysis. The alanine dehydrogenase from Vibrio proteolyticus and the diaminopimelate dehydrogenase from Symbiobacterium thermophilum were selected for in vitro production of (R)-3-fluoroalanine and (S)-3-fluoroalanine, respectively, using 3-fluoropyruvate as the substrate. Additionally, we discovered that an alanine racemase from Streptomyces lavendulae, originally selected for setting an alternative enzymatic cascade leading to the production of these non-canonical amino acids, had an unprecedented catalytic efficiency in β-elimination of fluorine from the monosubstituted fluoroalanine. The in vitro enzymatic cascade based on the dehydrogenases of V. proteolyticus and S. thermophilum included a cofactor recycling system, whereby a formate dehydrogenase from Pseudomonas sp. 101 (either native or engineered) coupled formate oxidation to NAD(P)H formation. Under these conditions, the reaction yields for (R)-3-fluoroalanine and (S)-3-fluoroalanine reached >85% on the fluorinated substrate and proceeded with complete enantiomeric excess. The selected dehydrogenases also catalyzed the conversion of trifluoropyruvate into trifluorinated alanine as a first-case example of fluorine biocatalysis with amino acids carrying a trifluoromethyl group.

Expanding the flexibility of base editing for high-throughput genetic screens in bacteria

Genome-wide screens have become powerful tools for elucidating genotype-to-phenotype relationships in bacteria. Of the varying techniques to achieve knockout and knockdown, CRISPR base editors are emerging as promising options. However, the limited number of available, efficient target sites hampers their use for high-throughput screening. Here, we make multiple advances to enable flexible base editing as part of high-throughput genetic screening in bacteria. We first co-opt the Streptococcus canis Cas9 that exhibits more flexible protospacer-adjacent motif recognition than the traditional Streptococcus pyogenes Cas9. We then expand beyond introducing premature stop codons by mutating start codons. Next, we derive guide design rules by applying machine learning to an essentiality screen conducted in Escherichia coli. Finally, we rescue poorly edited sites by combining base editing with Cas9-induced cleavage of unedited cells, thereby enriching for intended edits. The efficiency of this dual system was validated through a conditional essentiality screen based on growth in minimal media. Overall, expanding the scope of genome-wide knockout screens with base editors could further facilitate the investigation of new gene functions and interactions in bacteria.

Enhanced chaotrope tolerance and (S)-2-hydroxypropiophenone production by recombinant Pseudomonas putida engineered with Pprl from Deinococcus radiodurans

Pseudomonas putida is a soil bacterium with multiple uses in fermentation and biotransformation processes. P. putida ATCC 12633 can biotransform benzaldehyde and other aldehydes into valuable α-hydroxyketones, such as (S)-2-hydroxypropiophenone. However, poor tolerance of this strain toward chaotropic aldehydes hampers efficient biotransformation processes. To circumvent this problem, we expressed the gene encoding the global regulator PprI from Deinococcus radiodurans, an inducer of pleiotropic proteins promoting DNA repair, in P. putida. Fine-tuned gene expression was achieved using an expression plasmid under the control of the LacIQ /Ptrc system, and the cross-protective role of PprI was assessed against multiple stress treatments. Moreover, the stress-tolerant P. putida strain was tested for 2-hydroxypropiophenone production using whole resting cells in the presence of relevant aldehyde substrates. P. putida cells harbouring the global transcriptional regulator exhibited high tolerance toward benzaldehyde, acetaldehyde, ethanol, butanol, NaCl, H2 O2 and thermal stress, thereby reflecting the multistress protection profile conferred by PprI. Additionally, the engineered cells converted aldehydes to 2-hydroxypropiophenone more efficiently than the parental P. putida strain. 2-Hydroxypropiophenone concentration reached 1.6 g L-1 upon a 3-h incubation under optimized conditions, at a cell concentration of 0.033 g wet cell weight mL-1 in the presence of 20 mM benzaldehyde and 600 mM acetaldehyde. Product yield and productivity were 0.74 g 2-HPP g-1 benzaldehyde and 0.089 g 2-HPP g cell dry weight-1  h-1 , respectively, 35% higher than the control experiments. Taken together, these results demonstrate that introducing PprI from D. radiodurans enhances chaotrope tolerance and 2-HPP production in P. putida ATCC 12633.

The pAblo·pCasso self-curing vector toolset for unconstrained cytidine and adenine base-editing in Gram-negative bacteria

A synthetic biology toolkit, exploiting clustered regularly interspaced short palindromic repeats (CRISPR) and modified CRISPR-associated protein (Cas) base-editors, was developed for genome engineering in Gram-negative bacteria. Both a cytidine base-editor (CBE) and an adenine base-editor (ABE) have been optimized for precise single-nucleotide modification of plasmid and genome targets. CBE comprises a cytidine deaminase conjugated to a Cas9 nickase from Streptococcus pyogenes (SpnCas9), resulting in C→T (or G→A) substitutions. Conversely, ABE consists of an adenine deaminase fused to SpnCas9 for A→G (or T→C) editing. Several nucleotide substitutions were achieved using these plasmid-borne base-editing systems and a novel protospacer adjacent motif (PAM)-relaxed SpnCas9 (SpRY) variant. Base-editing was validated in Pseudomonas putida and other Gram-negative bacteria by inserting premature STOP codons into target genes, thereby inactivating both fluorescent proteins and metabolic (antibiotic-resistance) functions. The functional knockouts obtained by engineering STOP codons via CBE were reverted to the wild-type genotype using ABE. Additionally, a series of induction-responsive vectors was developed to facilitate the curing of the base-editing platform in a single cultivation step, simplifying complex strain engineering programs without relying on homologous recombination and yielding plasmid-free, modified bacterial cells.

Engineering extracellular electron transfer pathways of electroactive microorganisms by synthetic biology for energy and chemicals production

The excessive consumption of fossil fuels causes massive emission of CO2, leading to climate deterioration and environmental pollution. The development of substitutes and sustainable energy sources to replace fossil fuels has become a worldwide priority. Bio-electrochemical systems (BESs), employing redox reactions of electroactive microorganisms (EAMs) on electrodes to achieve a meritorious combination of biocatalysis and electrocatalysis, provide a green and sustainable alternative approach for bioremediation, CO2 fixation, and energy and chemicals production. EAMs, including exoelectrogens and electrotrophs, perform extracellular electron transfer (EET) (i.e., outward and inward EET), respectively, to exchange energy with the environment, whose rate determines the efficiency and performance of BESs. Therefore, we review the synthetic biology strategies developed in the last decade for engineering EAMs to enhance the EET rate in cell-electrode interfaces for facilitating the production of electricity energy and value-added chemicals, which include (1) progress in genetic manipulation and editing tools to achieve the efficient regulation of gene expression, knockout, and knockdown of EAMs; (2) synthetic biological engineering strategies to enhance the outward EET of exoelectrogens to anodes for electricity power production and anodic electro-fermentation (AEF) for chemicals production, including (i) broadening and strengthening substrate utilization, (ii) increasing the intracellular releasable reducing equivalents, (iii) optimizing c-type cytochrome (c-Cyts) expression and maturation, (iv) enhancing conductive nanowire biosynthesis and modification, (v) promoting electron shuttle biosynthesis, secretion, and immobilization, (vi) engineering global regulators to promote EET rate, (vii) facilitating biofilm formation, and (viii) constructing cell-material hybrids; (3) the mechanisms of inward EET, CO2 fixation pathway, and engineering strategies for improving the inward EET of electrotrophic cells for CO2 reduction and chemical production, including (i) programming metabolic pathways of electrotrophs, (ii) rewiring bioelectrical circuits for enhancing inward EET, and (iii) constructing microbial (photo)electrosynthesis by cell-material hybridization; (4) perspectives on future challenges and opportunities for engineering EET to develop highly efficient BESs for sustainable energy and chemical production. We expect that this review will provide a theoretical basis for the future development of BESs in energy harvesting, CO2 fixation, and chemical synthesis.

Deazaflavin metabolite produced by endosymbiotic bacteria controls fungal host reproduction

The endosymbiosis between the pathogenic fungus Rhizopus microsporus and the toxin-producing bacterium Mycetohabitans rhizoxinica represents a unique example of host control by an endosymbiont. Fungal sporulation strictly depends on the presence of endosymbionts as well as bacterially produced secondary metabolites. However, an influence of primary metabolites on host control remained unexplored. Recently, we discovered that M. rhizoxinica produces FO and 3PG-F420, a derivative of the specialized redox cofactor F420. Whether FO/3PG-F420 plays a role in the symbiosis has yet to be investigated. Here, we report that FO, the precursor of 3PG-F420, is essential to the establishment of a stable symbiosis. Bioinformatic analysis revealed that the genetic inventory to produce cofactor 3PG-F420 is conserved in the genomes of eight endofungal Mycetohabitans strains. By developing a CRISPR/Cas-assisted base editing strategy for M. rhizoxinica, we generated mutant strains deficient in 3PG-F420 (M. rhizoxinica ΔcofC) and in both FO and 3PG-F420 (M. rhizoxinica ΔfbiC). Co-culture experiments demonstrated that the sporulating phenotype of apo-symbiotic R. microsporus is maintained upon reinfection with wild-type M. rhizoxinica or M. rhizoxinica ΔcofC. In contrast, R. microsporus is unable to sporulate when co-cultivated with M. rhizoxinica ΔfbiC, even though the fungus was observed by super-resolution fluorescence microscopy to be successfully colonized. Genetic and chemical complementation of the FO deficiency of M. rhizoxinica ΔfbiC led to restoration of fungal sporulation, signifying that FO is indispensable for establishing a functional symbiosis. Even though FO is known for its light-harvesting properties, our data illustrate an important role of FO in inter-kingdom communication.

An expanded CRISPR-Cas9-assisted recombineering toolkit for engineering genetically intractable Pseudomonas aeruginosa isolates

Much of our current understanding of microbiology is based on the application of genetic engineering procedures. Since their inception (more than 30 years ago), methods based largely on allelic exchange and two-step selection processes have become a cornerstone of contemporary bacterial genetics. While these tools are established for adapted laboratory strains, they have limited applicability in clinical or environmental isolates displaying a large and unknown genetic repertoire that are recalcitrant to genetic modifications. Hence, new tools allowing genetic engineering of intractable bacteria must be developed to gain a comprehensive understanding of them in the context of their biological niche. Herein, we present a method for precise, efficient and rapid engineering of the opportunistic pathogen Pseudomonas aeruginosa. This procedure relies on recombination of short single-stranded DNA facilitated by targeted double-strand DNA breaks mediated by a synthetic Cas9 coupled with the efficient Ssr recombinase. Possible applications include introducing single-nucleotide polymorphisms, short or long deletions, and short DNA insertions using synthetic single-stranded DNA templates, drastically reducing the need of PCR and cloning steps. Our toolkit is encoded on two plasmids, harboring an array of different antibiotic resistance cassettes; hence, this approach can be successfully applied to isolates displaying natural antibiotic resistances. Overall, this toolkit substantially reduces the time required to introduce a range of genetic manipulations to a minimum of five experimental days, and enables a variety of research and biotechnological applications in both laboratory strains and difficult-to-manipulate P. aeruginosa isolates.

Recursive genome engineering decodes the evolutionary origin of an essential thymidylate kinase activity in Pseudomonas putida KT2440

IMPORTANCE

Investigating fundamental aspects of metabolism is vital for advancing our understanding of the diverse biochemical capabilities and biotechnological applications of bacteria. The origin of the essential thymidylate kinase function in the model bacterium Pseudomonas putida KT2440, seemingly interrupted due to the presence of a large genomic island that disrupts the cognate gene, eluded a satisfactory explanation thus far. This is a first-case example of an essential metabolic function, likely acquired by horizontal gene transfer, which "landed" in a locus encoding the same activity. As such, foreign DNA encoding an essential dNMPK could immediately adjust to the recipient host-instead of long-term accommodation and adaptation. Understanding how these functions evolve is a major biological question, and the work presented here is a decisive step toward this direction. Furthermore, identifying essential and accessory genes facilitates removing those deemed irrelevant in industrial settings-yielding genome-reduced cell factories with enhanced properties and genetic stability.

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.

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.

Precise genome engineering in Pseudomonas using phage-encoded homologous recombination and the Cascade-Cas3 system

A lack of generic and effective genetic manipulation methods for Pseudomonas has restricted fundamental research and utilization of this genus for biotechnology applications. Phage-encoded homologous recombination (PEHR) is an efficient tool for bacterial genome engineering. This PEHR system is based on a lambda Red-like operon (BAS) from Pseudomonas aeruginosa phage Ab31 and a Rac bacteriophage RecET-like operon (Rec-TEPsy) from P. syringae pv. syringae B728a and also contains exogenous elements, including the RecBCD inhibitor (Redγ or Pluγ) or single-stranded DNA-binding protein (SSB), that were added to enhance the PEHR recombineering efficiency. To solve the problem of false positives in Pseudomonas editing with the PEHR system, the processive enzyme Cas3 with a minimal Type I-C Cascade-based system was combined with PEHR. This protocol describes the utilization of a Pseudomonas-specific PEHR-Cas3 system that was designed to universally and proficiently modify the genomes of Pseudomonas species. The pipeline uses standardized cassettes combined with the concerted use of SacB counterselection and Cre site-specific recombinase for markerless or seamless genome modification, in association with vectors that possess the selectively replicating template R6K to minimize recombineering background. Compared with the traditional allelic exchange editing method, the PEHR-Cas3 system does not need to construct suicide plasmids carrying long homologous arms, thus simplifying the experimental procedure and shortening the traceless editing period. Compared with general editing systems based on phage recombinases, the PEHR-Cas3 system can effectively improve the screening efficiency of mutants using the cutting ability of Cas3 protein. The entire procedure requires ~12 days.

Systems Analysis of Highly Multiplexed CRISPR-Base Editing in Streptomycetes

CRISPR tools, especially Cas9n-sgRNA guided cytidine deaminase base editors such as CRISPR-BEST, have dramatically simplified genetic manipulation of streptomycetes. One major advantage of CRISPR base editing technology is the possibility to multiplex experiments in genomically instable species. Here, we demonstrate scaled up Csy4 based multiplexed genome editing using CRISPR-mcBEST in Streptomyces coelicolor. We evaluated the system by simultaneously targeting 9, 18, and finally all 28 predicted specialized metabolite biosynthetic gene clusters in a single experiment. We present important insights into the performance of Csy4 based multiplexed genome editing at different scales. Using multiomics analysis, we investigated the systems wide effects of such extensive editing experiments and revealed great potentials and important bottlenecks of CRISPR-mcBEST. The presented analysis provides crucial data and insights toward the development of multiplexed base editing as a novel paradigm for high throughput engineering of Streptomyces chassis and beyond.

Simultaneous multi-site editing of individual genomes using retron arrays

Our understanding of genomics is limited by the scale of our genomic technologies. While libraries of genomic manipulations scaffolded on CRISPR gRNAs have been transformative, these existing approaches are typically multiplexed across genomes. Yet much of the complexity of real genomes is encoded within a genome across sites. Unfortunately, building cells with multiple, non-adjacent precise mutations remains a laborious cycle of editing, isolating an edited cell, and editing again. Here, we describe a technology for precisely modifying multiple sites on a single genome simultaneously. This technology - termed a multitron - is built from a heavily modified retron, in which multiple donor-encoding msds are produced from a single transcript. The multitron architecture is compatible with both recombineering in prokaryotic cells and CRISPR editing in eukaryotic cells. We demonstrate applications for this approach in molecular recording, genetic element minimization, and metabolic engineering.

Highly Efficient Biosynthesis of Protocatechuic Acid via Recombinant Pseudomonas putida KT2440

Owing to their physiological activities, plant-derived phenolic acids, such as protocatechuic acid (PCA), have extensive applications and market prospects. However, traditional production processes present numerous challenges and cannot meet increasing market demands. Hence, we aimed to biosynthesize PCA by constructing an efficient microbial factory via metabolic engineering of Pseudomonas putida KT2440. Glucose metabolism was engineered by deleting the genes for gluconate 2-dehydrogenase to enhance PCA biosynthesis. To increase the biosynthetic metabolic flux, one extra copy of the genes aroG^opt, aroQ, and aroB was inserted into the genome. The resultant strain, KGVA04, produced 7.2 g/L PCA. By inserting the degradation tags GSD and DAS to decrease the amount of shikimate dehydrogenase, PCA biosynthesis was increased to 13.2 g/L in shake-flask fermentation and 38.8 g/L in fed-batch fermentation. To the best of our knowledge, this was the first use of degradation tags to adjust the amount of a key enzyme at the protein level in P. putida KT2440, evidencing the remarkable potential of this method for naturally producing phenolic acids.

Revolutionizing biofuel generation: Unleashing the power of CRISPR-Cas mediated gene editing of extremophiles

Molecular biology techniques like gene editing have altered the specific genes in micro-organisms to increase their efficiency to produce biofuels. This review paper investigates the outcomes of Clustered regularly interspaced short palindromic repeats (CRISPR) for gene editing in extremophilic micro-organisms to produce biofuel. Commercial production of biofuel from lignocellulosic waste is limited due to various constraints. A potential strategy to enhance the capability of extremophiles to produce biofuel is gene-editing via CRISPR-Cas technology. The efficiency of intracellular enzymes like cellulase, hemicellulose in extremophilic bacteria, fungi and microalgae has been increased by alteration of genes associated with enzymatic activity and thermotolerance. extremophilic microbes like Thermococcus kodakarensis, Thermotoga maritima, Thermus thermophilus, Pyrococcus furiosus and Sulfolobus sp. are explored for biofuel production. The conversion of lignocellulosic biomass into biofuels involves pretreatment, hydrolysis and fermentation. The challenges like off-target effect associated with use of extremophiles for biofuel production is also addressed. The appropriate regulations are required to maximize effectiveness while minimizing off-target cleavage, as well as the total biosafety of this technique. The latest discovery of the CRISPR-Cas system should provide a new channel in the creation of microbial biorefineries through site- specific gene editing that might boost the generation of biofuels from extremophiles. Overall, this review study highlights the potential for genome editing methods to improve the potential of extremophiles to produce biofuel, opening the door to more effective and environmentally friendly biofuel production methods.

Powerful Microbial Base-Editing Toolbox: From Optimization Strategies to Versatile Applications

Base editors (BE) based on CRISPR systems are practical gene-editing tools which continue to drive frontier advances of life sciences. BEs are able to efficiently induce point mutations at target sites without double-stranded DNA cleavage. Hence, they are widely employed in the fields of microbial genome engineering. As applications of BEs continue to expand, the demands for base-editing efficiency, fidelity, and versatility are also on the rise. In recent years, a series of optimization strategies for BEs have been developed. By engineering the core components of BEs or adopting different assembly methods, the performance of BEs has been well optimized. Moreover, series of newly established BEs have significantly expanded the base-editing toolsets. In this Review, we will summarize the current efforts for BE optimization, introduce several novel BEs with versatility, and look forward to the broadened applications for industrial microorganisms.

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.

Growth-coupled enzyme engineering through manipulation of redox cofactor regeneration

Enzymes need to be efficient, robust, and highly specific for their effective use in commercial bioproduction. These properties can be introduced using various enzyme engineering techniques, with random mutagenesis and directed evolution (DE) often being chosen when there is a lack of structural information -or mechanistic understanding- of the enzyme. The screening or selection step of DE is the limiting part of this process, since it must ideally be (ultra)-high throughput, specifically target the catalytic activity of the enzyme and have an accurately quantifiable metric for said activity. Growth-coupling selection strategies involve coupling a desired enzyme activity to cellular metabolism and therefore growth, where growth (rate) becomes the output metric. Redox cofactors (NAD⁺/NADH and NADP⁺/NADPH) have recently been identified as promising target molecules for growth coupling, owing to their essentiality for cellular metabolism and ubiquitous nature. Redox cofactor oxidation or reduction can be disrupted through metabolic engineering and the use of specific culturing conditions, rendering the cell inviable unless a 'rescue' reaction complements the imposed metabolic deficiency. Using this principle, enzyme variants displaying improved cofactor oxidation or reduction rates can be selected for through an increased growth rate of the cell. In recent years, several E. coli strains have been developed that are deficient in the oxidation or reduction of NAD⁺/NADH and NADP⁺/NADPH pairs, and of non-canonical redox cofactor pairs NMN⁺/NMNH and NCD⁺/NCDH, which provides researchers with a versatile toolbox of enzyme engineering platforms. A range of redox cofactor dependent enzymes have since been engineered using a variety of these strains, demonstrating the power of using this growth-coupling technique for enzyme engineering. This review aims to summarize the metabolic engineering involved in creating strains auxotrophic for the reduced or oxidized state of redox cofactors, and the resulting successes in using them for enzyme engineering. Perspectives on the unique features and potential future applications of this technique are also presented.

Bio-SCAN V2: A CRISPR/dCas9-based lateral flow assay for rapid detection of theophylline

Rapid, specific, and robust diagnostic strategies are needed to develop sensitive biosensors for small molecule detection, which could aid in controlling contamination and disease transmission. Recently, the target-induced collateral activity of Cas nucleases [clustered regularly interspaced short palindromic repeats (CRISPR)-associated nucleases] was exploited to develop high-throughput diagnostic modules for detecting nucleic acids and small molecules. Here, we have expanded the diagnostic ability of the CRISPR-Cas system by developing Bio-SCAN V2, a ligand-responsive CRISPR-Cas platform for detecting non-nucleic acid small molecule targets. The Bio-SCAN V2 consists of an engineered ligand-responsive sgRNA (ligRNA), biotinylated dead Cas9 (dCas9-biotin), 6-carboxyfluorescein (FAM)-labeled amplicons, and lateral flow assay (LFA) strips. LigRNA interacts with dCas9-biotin only in the presence of sgRNA-specific ligand molecules to make a ribonucleoprotein (RNP). Next, the ligand-induced ribonucleoprotein is exposed to FAM-labeled amplicons for binding, and the presence of the ligand (small molecule) is detected as a visual signal [(dCas9-biotin)-ligRNA-FAM labeled DNA-AuNP complex] at the test line of the lateral flow assay strip. With the Bio-SCAN V2 platform, we are able to detect the model molecule theophylline with a limit of detection (LOD) up to 2 μM in a short time, requiring only 15 min from sample application to visual readout. Taken together, Bio-SCAN V2 assay provides a rapid, specific, and ultrasensitive detection platform for theophylline.

Challenges and opportunities in bringing nonbiological atoms to life with synthetic metabolism

The relatively narrow spectrum of chemical elements within the microbial 'biochemical palate' limits the reach of biotechnology, because several added-value compounds can only be produced with traditional organic chemistry. Synthetic biology offers enabling tools to tackle this issue by facilitating 'biologization' of non-canonical chemical atoms. The interplay between xenobiology and synthetic metabolism multiplies routes for incorporating nonbiological atoms into engineered microbes. In this review, we survey natural assimilation routes for elements beyond the essential biology atoms [i.e., carbon (C), hydrogen (H), nitrogen (N), oxygen (O), phosphorus (P), and sulfur (S)], discussing how these mechanisms could be repurposed for biotechnology. Furthermore, we propose a computational framework to identify chemical elements amenable to biologization, ranking reactions suitable to build synthetic metabolism. When combined and deployed in robust microbial hosts, these approaches will offer sustainable alternatives for smart chemical production.

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 β-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.

Base editing for reprogramming cyanobacterium Synechococcus elongatus

Cyanobacteria can directly convert carbon dioxide (CO₂) at the atmospheric level to biofuels, value-added chemicals and food products, making them ideal candidates to alleviate global climate change. Despite decades-long pioneering successes, the development of genome-editing tools, especially the CRISPR-Cas-based approaches, seems to lag behind other microbial chassis, slowing down the innovations of cyanobacteria. Here, we adapted and tailored base editing for cyanobacteria based on the CRISPR-Cas system and deamination. We achieved precise and efficient genome editing at a single-nucleotide resolution and demonstrated multiplex base editing in the model cyanobacterium Synechococcus elongatus. By using the base-editing tool, we successfully manipulated the glycogen metabolic pathway via the introduction of premature STOP codons in the relevant genes, building engineered strains with elevated potentials to produce chemicals and food from CO₂. We present here the first report of base editing in the phylum of cyanobacteria, and a paradigm for applying CRISPR-Cas systems in bacteria. We believe that our work will accelerate the metabolic engineering and synthetic biology of cyanobacteria and drive more innovations to alleviate global climate change.

Synthetic metabolism for in vitro acetone biosynthesis driven by ATP regeneration

In vitro ketone production continues to be a challenge due to the biochemical features of the enzymes involved-even when some of them have been extensively characterized (e.g. thiolase from Clostridium acetobutylicum), the assembly of synthetic enzyme cascades still face significant limitations (including issues with protein aggregation and multimerization). Here, we designed and assembled a self-sustaining enzyme cascade with acetone yields close to the theoretical maximum using acetate as the only carbon input. The efficiency of this system was further boosted by coupling the enzymatic sequence to a two-step ATP-regeneration system that enables continuous, cost-effective acetone biosynthesis. Furthermore, simple methods were implemented for purifying the enzymes necessary for this synthetic metabolism, including a first-case example on the isolation of a heterotetrameric acetate:coenzyme A transferase by affinity chromatography.

Role of the CrcB transporter of Pseudomonas putida in the multi-level stress response elicited by mineral fluoride

The presence of mineral fluoride (F^- ) in the environment has both a geogenic and anthropogenic origin, and the halide has been described to be toxic in virtually all living organisms. While the evidence gathered in different microbial species supports this notion, a systematic exploration of the effects of F^- salts on the metabolism and physiology of environmental bacteria remained underexplored thus far. In this work, we studied and characterized tolerance mechanisms deployed by the model soil bacterium Pseudomonas putida KT2440 against NaF. By adopting systems-level omic approaches, including functional genomics and metabolomics, we gauged the impact of this anion at different regulatory levels under conditions that impair bacterial growth. Several genes involved in halide tolerance were isolated in a genome-wide Tn-Seq screening-among which crcB, encoding an F^- -specific exporter, was shown to play the predominant role in detoxification. High-resolution metabolomics, combined with the assessment of intracellular and extracellular pH values and quantitative physiology experiments, underscored the key nodes in central carbon metabolism affected by the presence of F^- . Taken together, our results indicate that P. putida undergoes a general, multi-level stress response when challenged with NaF that significantly differs from that caused by other saline stressors. While microbial stress responses to saline and oxidative challenges have been extensively studied and described in the literature, very little is known about the impact of fluoride (F^- ) on bacterial physiology and metabolism. This state of affairs contrasts with the fact that F^- is more abundant than other halides in the Earth crust (e.g. in some soils, the F^- concentration can reach up to 1 mg g_soil ^-1 ). Understanding the global effects of NaF treatment on bacterial physiology is not only relevant to unveil distinct mechanisms of detoxification but it could also guide microbial engineering approaches for the target incorporation of fluorine into value-added organofluorine molecules. In this regard, the soil bacterium P. putida constitutes an ideal model to explore such scenarios, since this species is particularly known for its high level of stress resistance against a variety of physicochemical perturbations.

Highly efficient multiplex base editing: One-shot deactivation of eight genes in Shewanella oneidensis MR-1

Obtaining electroactive microbes capable of efficient extracellular electron transfer is a large undertaking for the scalability of bio-electrochemical systems. Inevitably, researchers need to pursue the co-modification of multiple genes rather than expecting that modification of a single gene would make a significant contribution to improving extracellular electron transfer rates. Base editing has enabled highly-efficient gene deactivation in model electroactive microbe Shewanella oneidensis MR-1. Since multiplexed application of base editing is still limited by its low throughput procedure, we thus here develop a rapid and efficient multiplex base editing system in S. oneidensis. Four approaches to express multiple gRNAs were assessed firstly, and transcription of each gRNA cassette into a monocistronic unit was validated as a more favorable option than transcription of multiple gRNAs into a polycistronic cluster. Then, a smart scheme was designed to deliver one-pot assembly of multiple gRNAs. 3, 5, and 8 genes were deactivated using this system with editing efficiency of 83.3%, 100% and 12.5%, respectively. To offer some nonrepetitive components as alternatives genetic parts of sgRNA cassette, different promoters, handles, and terminators were screened. This multiplex base editing tool was finally adopted to simultaneously deactivate eight genes that were identified as significantly downregulated targets in transcriptome analysis of riboflavin-overproducing strain and control strain. The maximum power density of the multiplex engineered strain HRF(8BE) in microbial fuel cells was 1108.1 mW/m², which was 21.67 times higher than that of the wild-type strain. This highly efficient multiplexed base editing tool elevates our ability of genome manipulation and combinatorial engineering in Shewanella, and may provide valuable insights in fundamental and applied research of extracellular electron transfer.

A synthetic C2 auxotroph of Pseudomonas putida for evolutionary engineering of alternative sugar catabolic routes

Acetyl-coenzyme A (AcCoA) is a metabolic hub in virtually all living cells, serving as both a key precursor of essential biomass components and a metabolic sink for catabolic pathways for a large variety of substrates. Owing to this dual role, tight growth-production coupling schemes can be implemented around the AcCoA node. Building on this concept, a synthetic C2 auxotrophy was implemented in the platform bacterium Pseudomonas putida through an in silico-informed engineering approach. A growth-coupling strategy, driven by AcCoA demand, allowed for direct selection of an alternative sugar assimilation route-the phosphoketolase (PKT) shunt from bifidobacteria. Adaptive laboratory evolution forced the synthetic P. putida auxotroph to rewire its metabolic network to restore C2 prototrophy via the PKT shunt. Large-scale structural chromosome rearrangements were identified as possible mechanisms for adjusting the network-wide proteome profile, resulting in improved PKT-dependent growth phenotypes. ¹³C-based metabolic flux analysis revealed an even split between the native Entner-Doudoroff pathway and the synthetic PKT bypass for glucose processing, leading to enhanced carbon conservation. These results demonstrate that the P. putida metabolism can be radically rewired to incorporate a synthetic C2 metabolism, creating novel network connectivities and highlighting the importance of unconventional engineering strategies to support efficient microbial production.