A highly precise and portable genome engineering method allows comparison of mutational effects across bacterial species

Improved dsDNA recombineering enables versatile multiplex genome engineering of kilobase-scale sequences in diverse bacteria

Recombineering assisted multiplex genome editing generally uses single-stranded oligonucleotides for site directed mutational changes. It has proven highly efficient for functional screens and to optimize microbial cell factories. However, this approach is limited to relatively small mutational changes. Here, we addressed the challenges involved in the use of double-stranded DNA substrates for multiplex genome engineering. Recombineering is mediated by phage single-strand annealing proteins annealing ssDNAs into the replication fork. We apply this insight to facilitate the generation of ssDNA from the dsDNA substrate and to alter the speed of replication by elevating the available deoxynucleoside triphosphate (dNTP) levels. Intracellular dNTP concentration was elevated by ribonucleotide reductase overexpression or dNTP addition to establish double-stranded DNA Recombineering-assisted Multiplex Genome Engineering (dReaMGE), which enables rapid and flexible insertional and deletional mutagenesis at multiple sites on kilobase scales in diverse bacteria without the generation of double-strand breaks or disturbance of the mismatch repair system. dReaMGE can achieve combinatorial genome engineering works, for example, alterations to multiple biosynthetic pathways, multiple promoter or gene insertions, variations of transcriptional regulator combinations, within a few days. dReaMGE adds to the repertoire of bacterial genome engineering to facilitate discovery, functional genomics, strain optimization and directed evolution of microbial cell factories.

Plasmid-mediated ciprofloxacin resistance imparts a selective advantage on Escherichia coli ST131

Escherichia coli ST131 is a recently emerged antibiotic resistant clone responsible for high rates of urinary tract and bloodstream infections. Despite its global dominance, the precise mechanisms that have driven the rapid dissemination of ST131 remain unknown. Here, we show that the plasmid-associated resistance gene encoding the AAC(6')-Ib-cr enzyme that inactivates the fluoroquinolone antibiotic ciprofloxacin is present in >70% of strains from the most rapidly expanding subgroup of multidrug resistant ST131. Using a series of genome-edited and plasmid-cured isogenic strains, we demonstrate that the aac(6')-Ib-cr gene confers a selective advantage on ST131 in the presence of ciprofloxacin, even in strains containing chromosomal GyrA and ParC FQ-resistance mutations. Further, we identify a pattern of emerging carbapenem resistance in other common E. coli clones carrying both aac(6')-Ib-cr and chromosomal FQ-resistance mutations, suggesting this dual resistance combination may also impart a selective advantage on these non-ST131 antibiotic resistant lineages.

Status quo of tet regulation in bacteria

The tetracycline repressor (TetR) belongs to the most popular, versatile and efficient transcriptional regulators used in bacterial genetics. In the tetracycline (Tc) resistance determinant tet(B) of transposon Tn10, tetR regulates the expression of a divergently oriented tetA gene that encodes a Tc antiporter. These components of Tn10 and of other natural or synthetic origins have been used for tetracycline‐dependent gene regulation (tet regulation) in at least 40 bacterial genera. Tet regulation serves several purposes such as conditional complementation, depletion of essential genes, modulation of artificial genetic networks, protein overexpression or the control of gene expression within cell culture or animal infection models. Adaptations of the promoters employed have increased tet regulation efficiency and have made this system accessible to taxonomically distant bacteria. Variations of TetR, different effector molecules and mutated DNA binding sites have enabled new modes of gene expression control. This article provides a current overview of tet regulation in bacteria.

Golden Gate Assembly of Aerobic and Anaerobic Microbial Bioreporters

Microbial bioreporters provide direct insight into cellular processes by producing a quantifiable signal dictated by reporter gene expression. The core of a bioreporter is a genetic circuit in which a reporter gene (or operon) is fused to promoter and regulatory sequences that govern its expression. In this study, we develop a system for constructing novel Escherichia coli bioreporters based on Golden Gate assembly, a synthetic biology approach for the rapid and seamless fusion of DNA fragments. Gene circuits are generated by fusing promoter and reporter sequences encoding yellow fluorescent protein, mCherry, bacterial luciferase, and an anaerobically active flavin-based fluorescent protein. We address a barrier to the implementation of Golden Gate assembly by designing a series of compatible destination vectors that can accommodate the assemblies. We validate the approach by measuring the activity of constitutive bioreporters and mercury and arsenic biosensors in quantitative exposure assays. We also demonstrate anaerobic quantification of mercury and arsenic in biosensors that produce flavin-based fluorescent protein, highlighting the expanding range of redox conditions that can be examined by microbial bioreporters. IMPORTANCE Microbial bioreporters are versatile genetic tools with wide-ranging applications, particularly in the field of environmental toxicology. For example, biosensors that produce a signal output in the presence of a specific analyte offer less costly alternatives to analytical methods for the detection of environmental toxins such as mercury and arsenic. Biosensors of specific toxins can also be used to test hypotheses regarding mechanisms of uptake, toxicity, and biotransformation. In this study, we develop an assembly platform that uses a synthetic biology technique to streamline construction of novel Escherichia coli bioreporters that produce fluorescent or luminescent signals either constitutively or in response to mercury and arsenic exposure. Beyond the synthesis of novel biosensors, our assembly platform can be adapted for numerous applications, including labelling bacteria for fluorescent microscopy, developing gene expression systems, and modifying bacterial genomes.

ReScribe: An Unrestrained Tool Combining Multiplex Recombineering and Minimal-PAM ScCas9 for Genome Recoding Pseudomonas putida

Genome recoding enables incorporating new functions into the DNA of microorganisms. By reassigning codons to noncanonical amino acids, the generation of new-to-nature proteins offers countless opportunities for bioproduction and biocontainment in industrial chassis. A key bottleneck in genome recoding efforts, however, is the low efficiency of recombineering, which hinders large-scale applications at acceptable speed and cost. To relieve this bottleneck, we developed ReScribe, a highly optimized recombineering tool enhanced by CRISPR-Cas9-mediated counterselection built upon the minimal PAM 5'-NNG-3' of the Streptococcus canis Cas9 (ScCas9). As a proof of concept, we used ReScribe to generate a minimally recoded strain of the industrial chassis Pseudomonas putida by replacing TAG stop codons (functioning as PAMs) of essential metabolic genes with the synonymous TAA. We showed that ReScribe enables nearly 100% engineering efficiency of multiple loci in P. putida, opening promising avenues for genome editing and applications thereof in this bacterium and beyond.

ssDNA recombineering boosts in vivo evolution of nanobodies displayed on bacterial surfaces

ssDNA recombineering has been exploited to hyperdiversify genomically-encoded nanobodies displayed on the surface of Escherichia coli for originating new binding properties. As a proof-of-principle a nanobody recognizing the antigen TirM from enterohaemorrhagic E. coli (EHEC) was evolved towards the otherwise not recognized TirM antigen from enteropathogenic E. coli (EPEC). To this end, E. coli cells displaying this nanobody fused to the intimin outer membrane-bound domain were subjected to multiple rounds of mutagenic oligonucleotide recombineering targeting the complementarity determining regions (CDRs) of the cognate VHH gene sequence. Binders to the EPEC-TirM were selected upon immunomagnetic capture of bacteria bearing active variants and nanobodies identified with a new ability to strongly bind the new antigen. The results highlight the power of combining evolutionary properties of bacteria in vivo with oligonucleotide synthesis in vitro for the sake of focusing diversification to specific segments of a gene (or protein thereof) of interest.

A standardized genome architecture for bacterial synthetic biology (SEGA)

Chromosomal recombinant gene expression offers a number of advantages over plasmid-based synthetic biology. However, the methods applied for bacterial genome engineering are still challenging and far from being standardized. Here, in an attempt to realize the simplest recombinant genome technology imaginable and facilitate the transition from recombinant plasmids to genomes, we create a simplistic methodology and a comprehensive strain collection called the Standardized Genome Architecture (SEGA). In its simplest form, SEGA enables genome engineering by combining only two reagents: a DNA fragment that can be ordered from a commercial vendor and a stock solution of bacterial cells followed by incubation on agar plates. Recombinant genomes are identified by visual inspection using green-white colony screening akin to classical blue-white screening for recombinant plasmids. The modular nature of SEGA allows precise multi-level control of transcriptional, translational, and post-translational regulation. The SEGA architecture simultaneously supports increased standardization of genetic designs and a broad application range by utilizing well-characterized parts optimized for robust performance in the context of the bacterial genome. Ultimately, its adaption and expansion by the scientific community should improve predictability and comparability of experimental outcomes across different laboratories.

Pseudomonas aeruginosa adapts to octenidine via a combination of efflux and membrane remodelling

Pseudomonas aeruginosa is an opportunistic pathogen capable of stably adapting to the antiseptic octenidine by an unknown mechanism. Here we characterise this adaptation, both in the laboratory and a simulated clinical setting, and identify a novel antiseptic resistance mechanism. In both settings, 2 to 4-fold increase in octenidine tolerance was associated with stable mutations and a specific 12 base pair deletion in a putative Tet-repressor family gene (smvR), associated with a constitutive increase in expression of the Major Facilitator Superfamily (MFS) efflux pump SmvA. Adaptation to higher octenidine concentrations led to additional stable mutations, most frequently in phosphatidylserine synthase pssA and occasionally in phosphatidylglycerophosphate synthase pgsA genes, resulting in octenidine tolerance 16- to 256-fold higher than parental strains. Metabolic changes were consistent with mitigation of oxidative stress and altered plasma membrane composition and order. Mutations in SmvAR and phospholipid synthases enable higher level, synergistic tolerance of octenidine.

Simultaneous Multiplex Genome Engineering via Accelerated Natural Transformation in Bacillus subtilis

Multiplex engineering at the scale of whole genomes has become increasingly important for synthetic biology and biotechnology applications. Although several methods have been reported for engineering microbe genomes, their use is limited by their complex procedures using multi-cycle transformations. Natural transformation, involving in species evolution by horizontal gene transfer in many organisms, indicates its potential as a genetic tool. Here, we aimed to develop simultaneous multiplex genome engineering (SMGE) for the simple, rapid, and efficient design of bacterial genomes via one-step of natural transformation in Bacillus subtilis. The transformed DNA, competency factors, and recombinases were adapted to improved co-editing frequencies above 27-fold. Single to octuplet variants with genetic diversity were simultaneously generated using all-in-one vectors harboring multi-gene cassettes. To demonstrate its potential application, the tyrosine biosynthesis pathway was further optimized for producing commercially important resveratrol by high-throughput screening of variant pool in B. subtilis. SMGE represents an accelerated evolution platform that generates diverse multiplex mutations for large-scale genetic engineering and synthetic biology in B. subtilis.

A versatile genetic engineering toolkit for E. coli based on CRISPR-prime editing

CRISPR base editing is a powerful method to engineer bacterial genomes. However, it restricts editing to single-nucleotide substitutions. Here, to address this challenge, we adapt a CRISPR-Prime Editing-based, DSB-free, versatile, and single-nucleotide resolution genetic manipulation toolkit for prokaryotes. It can introduce substitutions, deletions, insertions, and the combination thereof, both in plasmids and the chromosome of E. coli with high fidelity. Notably, under optimal conditions, the efficiency of 1-bp deletions reach up to 40%. Moreover, deletions of up to 97 bp and insertions up to 33 bp were successful with the toolkit in E. coli, however, efficiencies dropped sharply with increased fragment sizes. With a second guide RNA, our toolkit can achieve multiplexed editing albeit with low efficiency. Here we report not only a useful addition to the genome engineering arsenal for E. coli, but also a potential basis for the development of similar toolkits for other bacteria.

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

The nicotinamide cofactor specificity of enzymes plays a key role in regulating metabolic processes and attaining cellular homeostasis. Multiple studies have used enzyme engineering tools or a directed evolution approach to switch the cofactor preference of specific oxidoreductases. However, whole-cell adaptation toward the emergence of novel cofactor regeneration routes has not been previously explored. To address this challenge, we used an Escherichia coli NADPH-auxotrophic strain. We continuously cultivated this strain under selective conditions. After 500 to 1,100 generations of adaptive evolution using different carbon sources, we isolated several strains capable of growing without an external NADPH source. Most isolated strains were found to harbor a mutated NAD⁺-dependent malic enzyme (MaeA). A single mutation in MaeA was found to switch cofactor specificity while lowering enzyme activity. Most mutated MaeA variants also harbored a second mutation that restored the catalytic efficiency of the enzyme. Remarkably, the best MaeA variants identified this way displayed overall superior kinetics relative to the wild-type variant with NAD⁺. In other evolved strains, the dihydrolipoamide dehydrogenase (Lpd) was mutated to accept NADP⁺, thus enabling the pyruvate dehydrogenase and 2-ketoglutarate dehydrogenase complexes to regenerate NADPH. Interestingly, no other central metabolism oxidoreductase seems to evolve toward reducing NADP⁺, which we attribute to several biochemical constraints, including unfavorable thermodynamics. This study demonstrates the potential and biochemical limits of evolving oxidoreductases within the cellular context toward changing cofactor specificity, further showing that long-term adaptive evolution can optimize enzyme activity beyond what is achievable via rational design or directed evolution using small libraries.

Directed Evolution: Methodologies and Applications

Directed evolution aims to expedite the natural evolution process of biological molecules and systems in a test tube through iterative rounds of gene diversifications and library screening/selection. It has become one of the most powerful and widespread tools for engineering improved or novel functions in proteins, metabolic pathways, and even whole genomes. This review describes the commonly used gene diversification strategies, screening/selection methods, and recently developed continuous evolution strategies for directed evolution. Moreover, we highlight some representative applications of directed evolution in engineering nucleic acids, proteins, pathways, genetic circuits, viruses, and whole cells. Finally, we discuss the challenges and future perspectives in directed evolution.

Genetic Diversity for Accelerating Microbial Adaptive Laboratory Evolution

Adaptive laboratory evolution (ALE) is a widely used and highly effective tool for improving microbial phenotypes and investigating the evolutionary roots of biological phenomena. Serving as the raw materials of evolution, mutations have been extensively utilized to increase the chances of engineering molecules or microbes with tailor-made functions. The generation of genetic diversity is therefore a core technology for accelerating ALE, and a high-quality mutant library is crucial to its success. Because of its importance, technologies for generating genetic diversity have undergone rapid development in recent years. Here, we review the existing techniques for the construction of mutant libraries, briefly introduce their mechanisms and applications, discuss ongoing and emerging efforts to apply engineering technologies in the construction of mutant libraries, and suggest future perspectives for library construction.

Analysing Parallel Strategies to Alter the Host Specificity of Bacteriophage T7

Simple Summary

The problem of antimicrobial resistance is prominent and new alternatives to antibiotics are necessary. Bacteriophages are viruses that target host bacteria and can be used efficiently for their antibacterial properties to solve the problem of antimicrobial resistance. In this study, we explore ways to genetically modify T7 bacteriophage and make its tropism broader, so that it can attack a higher variety of bacteria. We are using different methodologies to achieve this, among of those bacteriophage recombineering using electroporated DNA (BRED), which seems to be the most efficient.

Abstract

The recognition and binding of host bacteria by bacteriophages is most often enabled by a highly specific receptor–ligand type of interaction, with the receptor-binding proteins (RBPs) of phages being the primary determinants of host specificity. Specifically modifying the RBPs could alter or extend the host range of phages otherwise exhibiting desired phenotypic properties. This study employed two different strategies to reprogram T7 phages ordinarily infecting commensal K12 Escherichia coli strains to infect pathogen-associated K1-capsule-expressing strains. The strategies were based on either plasmid-based homologous recombination or bacteriophage recombineering using electroporated DNA (BRED). Our work pursued the construction of two genetic designs: one replacing the gp17 gene of T7, the other replacing gp11, gp12, and gp17 of T7 with their K1F counterparts. Both strategies displayed successful integration of the K1F sequences into the T7 genome, detected by PCR screening. Multiple methods were utilised to select or enrich for chimeric phages incorporating the K1F gp17 alone, including trxA, host-specificity, and CRISPR-Cas-based selection. Irrespective of the selection method, the above strategy yielded poorly reproducible phage propagation on the new host, indicating that the chimeric phage was less fit than the wild type and could not promote continual autonomous reproduction. Chimeric phages obtained from BRED incorporating gp11-12 and gp17, however, all displayed infection in a 2-stage pattern, indicating the presence of both K1F and T7 phenotypes. This study shows that BRED can be used as a tool to quickly access the potential of new RBP constructs without the need to engineer sustainably replicating phages. Additionally, we show that solely repurposing the primary RBP is, in some cases, insufficient to produce a viable chimeric phage.

Spatiotemporal Manipulation of the Mismatch Repair System of Pseudomonas putida Accelerates Phenotype Emergence

The development of complex phenotypes in industrially relevant bacteria is a major goal of metabolic engineering, which encompasses the implementation of both rational and random approaches. In the latter case, several tools have been developed toward increasing mutation frequencies, yet the precise control of mutagenesis processes in cell factories continues to represent a significant technical challenge. Pseudomonas species are endowed with one of the most efficient DNA mismatch repair (MMR) systems found in the bacterial domain. Here, we investigated if the endogenous MMR system could be manipulated as a general strategy to artificially alter mutation rates in Pseudomonas species. To bestow a conditional mutator phenotype in the platform bacterium Pseudomonas putida, we constructed inducible mutator devices to modulate the expression of the dominant-negative mutL^E36K allele. Regulatable overexpression of mutL^E36K in a broad-host-range, easy-to-cure plasmid format resulted in a transitory inhibition of the MMR machinery, leading to a significant increase (up to 438-fold) in DNA mutation frequencies and a heritable fixation of mutations in the genome. Following such an accelerated mutagenesis-followed by selection approach, three phenotypes were successfully evolved: resistance to antibiotics streptomycin and rifampicin (either individually or combined) and reversion of a synthetic uracil auxotrophy. Thus, these mutator devices could be applied to accelerate the evolution of metabolic pathways in long-term evolutionary experiments, alternating cycles of (inducible) mutagenesis coupled to selection schemes toward the desired phenotype(s).

High-throughput functional variant screens via in vivo production of single-stranded DNA

Creating and characterizing individual genetic variants remains limited in scale, compared to the tremendous variation both existing in nature and envisioned by genome engineers. Here we introduce retron library recombineering (RLR), a methodology for high-throughput functional screens that surpasses the scale and specificity of CRISPR-Cas methods. We use the targeted reverse-transcription activity of retrons to produce single-stranded DNA (ssDNA) in vivo, incorporating edits at >90% efficiency and enabling multiplexed applications. RLR simultaneously introduces many genomic variants, producing pooled and barcoded variant libraries addressable by targeted deep sequencing. We use RLR for pooled phenotyping of synthesized antibiotic resistance alleles, demonstrating quantitative measurement of relative growth rates. We also perform RLR using the sheared genomic DNA of an evolved bacterium, experimentally querying millions of sequences for causal variants, demonstrating that RLR is uniquely suited to utilize large pools of natural variation. Using ssDNA produced in vivo for pooled experiments presents avenues for exploring variation across the genome.

Characterizing the portability of phage-encoded homologous recombination proteins

Efficient genome editing methods are essential for biotechnology and fundamental research. Homologous recombination (HR) is the most versatile method of genome editing, but techniques that rely on host RecA-mediated pathways are inefficient and laborious. Phage-encoded single-stranded DNA annealing proteins (SSAPs) improve HR 1,000-fold above endogenous levels. However, they are not broadly functional. Using Escherichia coli, Lactococcus lactis, Mycobacterium smegmatis, Lactobacillus rhamnosus and Caulobacter crescentus, we investigated the limited portability of SSAPs. We find that these proteins specifically recognize the C-terminal tail of the host's single-stranded DNA-binding protein (SSB) and are portable between species only if compatibility with this host domain is maintained. Furthermore, we find that co-expressing SSAPs with SSBs can significantly improve genome editing efficiency, in some species enabling SSAP functionality even without host compatibility. Finally, we find that high-efficiency HR far surpasses the mutational capacity of commonly used random mutagenesis methods, generating exceptional phenotypes that are inaccessible through sequential nucleotide conversions.

A navigation guide of synthetic biology tools for Pseudomonas putida

Pseudomonas putida is a microbial chassis of huge potential for industrial and environmental biotechnology, owing to its remarkable metabolic versatility and ability to sustain difficult redox reactions and operational stresses, among other attractive characteristics. A wealth of genetic and in silico tools have been developed to enable the unravelling of its physiology and improvement of its performance. However, the rise of this microbe as a promising platform for biotechnological applications has resulted in diversification of tools and methods rather than standardization and convergence. As a consequence, multiple tools for the same purpose have been generated, whilst most of them have not been embraced by the scientific community, which has led to compartmentalization and inefficient use of resources. Inspired by this and by the substantial increase in popularity of P. putida, we aim herein to bring together and assess all currently available (wet and dry) synthetic biology tools specific for this microbe, focusing on the last 5 years. We provide information on the principles, functionality, advantages and limitations, with special focus on their use in metabolic engineering. Additionally, we compare the tool portfolio for P. putida with those for other bacterial chassis and discuss potential future directions for tool development. Therefore, this review is intended as a reference guide for experts and new 'users' of this promising chassis.

Optimizing recombineering in Corynebacterium glutamicum

Owing to the increasing demand for amino acids and valuable commodities that can be produced by Corynebacterium glutamicum, there is a pressing need for new rapid genome engineering tools that improve the speed and efficiency of genomic insertions, deletions, and mutations. Recombineering using the λ Red system in Escherichia coli has proven very successful at genetically modifying this organism in a quick and efficient manner, suggesting that optimizing a recombineering system for C. glutamicum will also improve the speed for genomic modifications. Here, we maximized the recombineering efficiency in C. glutamicum by testing the efficacy of seven different recombinase/exonuclease pairs for integrating single-stranded DNA and double-stranded DNA (dsDNA) into the genome. By optimizing the homologous arm length and the amount of dsDNA transformed, as well as eliminating codon bias, a dsDNA recombineering efficiency of 13,250 transformed colonies/10⁹ viable cells was achieved, the highest efficiency currently reported in the literature. Using this optimized system, over 40,000 bp could be deleted in one transformation step. This recombineering strategy will greatly improve the speed of genetic modifications in C. glutamicum and assist other systems, such as clustered regularly interspaced short palindromic repeats and multiplexed automated genome engineering, in improving targeted genome editing.

Substrate-engaged type III secretion system structures reveal gating mechanism for unfolded protein translocation

Many bacterial pathogens rely on virulent type III secretion systems (T3SSs) or injectisomes to translocate effector proteins in order to establish infection. The central component of the injectisome is the needle complex which assembles a continuous conduit crossing the bacterial envelope and the host cell membrane to mediate effector protein translocation. However, the molecular principles underlying type III secretion remain elusive. Here, we report a structure of an active Salmonella enterica serovar Typhimurium needle complex engaged with the effector protein SptP in two functional states, revealing the complete 800Å-long secretion conduit and unraveling the critical role of the export apparatus (EA) subcomplex in type III secretion. Unfolded substrates enter the EA through a hydrophilic constriction formed by SpaQ proteins, which enables side chain-independent substrate transport. Above, a methionine gasket formed by SpaP proteins functions as a gate that dilates to accommodate substrates while preventing leaky pore formation. Following gate penetration, a moveable SpaR loop first folds up to then support substrate transport. Together, these findings establish the molecular basis for substrate translocation through T3SSs and improve our understanding of bacterial pathogenicity and motility.

A lysate proteome engineering strategy for enhancing cell-free metabolite production

Cell-free systems present a significant opportunity to harness the metabolic potential of diverse organisms. Removing the cellular context provides the ability to produce biological products without the need to maintain cell viability and enables metabolic engineers to explore novel chemical transformation systems. Crude extracts maintain much of a cell’s capabilities. However, only limited tools are available for engineering the contents of the extracts used for cell-free systems. Thus, our ability to take full advantage of the potential of crude extracts for cell-free metabolic engineering is constrained. Here, we employ Multiplex Automated Genomic Engineering (MAGE) to tag proteins for selective depletion from crude extracts so as to specifically direct chemical production. Specific edits to central metabolism are possible without significantly impacting cell growth. Selective removal of pyruvate degrading enzymes resulted in engineered crude lysates that are capable of up to 40-fold increases in pyruvate production when compared to the non-engineered extract. The described approach melds the tools of systems and synthetic biology to showcase the effectiveness of cell-free metabolic engineering for applications like bioprototyping and bioproduction.

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A method of engineering cell-free metabolism in lysates is described.

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Method enables design of cell lysates for enhancing specific metabolic processes.

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Pyruvate consuming enzymes tagged with 6xHis tags have minimal impact on growth.

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Post-lysis pull-down of tagged enzymes enables cell-free pyruvate pooling.

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Lysate engineering strategy permits metabolic states not possible in living cells.

New dual ATP-competitive inhibitors of bacterial DNA gyrase and topoisomerase IV active against ESKAPE pathogens

The rise in multidrug-resistant bacteria defines the need for identification of new antibacterial agents that are less prone to resistance acquisition. Compounds that simultaneously inhibit multiple bacterial targets are more likely to suppress the evolution of target-based resistance than monotargeting compounds. The structurally similar ATP binding sites of DNA gyrase and topoisomerase Ⅳ offer an opportunity to accomplish this goal. Here we present the design and structure-activity relationship analysis of balanced, low nanomolar inhibitors of bacterial DNA gyrase and topoisomerase IV that show potent antibacterial activities against the ESKAPE pathogens. For inhibitor 31c, a crystal structure in complex with Staphylococcus aureus DNA gyrase B was obtained that confirms the mode of action of these compounds. The best inhibitor, 31h, does not show any in vitro cytotoxicity and has excellent potency against Gram-positive (MICs: range, 0.0078-0.0625 μg/mL) and Gram-negative pathogens (MICs: range, 1-2 μg/mL). Furthermore, 31h inhibits GyrB mutants that can develop resistance to other drugs. Based on these data, we expect that structural derivatives of 31h will represent a step toward clinically efficacious multitargeting antimicrobials that are not impacted by existing antimicrobial resistance.

Engineering Bacteria to Produce Pure Phage-like Particles for Gene Delivery

Natural and engineered phages have been used in many applications, but their use to deliver user-defined genetic cargoes has been hampered by contamination with replicative phage, restricting use of the technology beyond the laboratory. Here we present a method to produce transducing particles without contamination. In addition, we demonstrate the use of a helper phage-free transducing particle preparation as an antimicrobial agent. This will pave the way for the development of new phage-based technologies with greater scope than lytic phage therapy.

Recombineering and MAGE

Recombination-mediated genetic engineering, also known as recombineering, is the genomic incorporation of homologous single-stranded or double-stranded DNA into bacterial genomes. Recombineering and its derivative methods have radically improved genome engineering capabilities, perhaps none more so than multiplex automated genome engineering (MAGE). MAGE is representative of a set of highly multiplexed single-stranded DNA-mediated technologies. First described in Escherichia coli, both MAGE and recombineering are being rapidly translated into diverse prokaryotes and even into eukaryotic cells. Together, this modern set of tools offers the promise of radically improving the scope and throughput of experimental biology by providing powerful new methods to ease the genetic manipulation of model and non-model organisms. In this Primer, we describe recombineering and MAGE, their optimal use, their diverse applications and methods for pairing them with other genetic editing tools. We then look forward to the future of genetic engineering.

In vivo diversification of target genomic sites using processive base deaminase fusions blocked by dCas9

In vivo mutagenesis systems accelerate directed protein evolution but often show restricted capabilities and deleterious off-site mutations on cells. To overcome these limitations, here we report an in vivo platform to diversify specific DNA segments based on protein fusions between various base deaminases (BD) and the T7 RNA polymerase (T7RNAP) that recognizes a cognate promoter oriented towards the target sequence. Transcriptional elongation of these fusions generates transitions C to T or A to G on both DNA strands and in long DNA segments. To delimit the boundaries of the diversified DNA, the catalytically dead Cas9 (dCas9) is tethered with custom-designed crRNAs as a "roadblock" for BD-T7RNAP elongation. Using this T7-targeted dCas9-limited in vivo mutagenesis (T7-DIVA) system, rapid molecular evolution of the antibiotic resistance gene TEM-1 is achieved. While the efficiency is demonstrated in E. coli, the system can be adapted to a variety of bacterial and eukaryotic hosts.

Emergence of the Novel Aminoglycoside Acetyltransferase Variant aac(6')-Ib-D179Y and Acquisition of Colistin Heteroresistance in Carbapenem-Resistant Klebsiella pneumoniae Due to a Disrupting Mutation in the DNA Repair Enzyme MutS

Amikacin and colistin are effective against carbapenem-resistant Klebsiella pneumoniae In 2017, we successively isolated three carbapenem-resistant K. pneumoniae isolates (ST967) from a patient with chronic renal failure in Japan. The first (SMKP01, sputum, day 0) and second (SMKP02, blood, day 14) strains were resistant to most antimicrobials tested but still susceptible to amikacin (MICs of 4 and 0.5 mg/liter, respectively) and colistin (MIC of 0.5 mg/liter for both). The third strain (SMKP03, blood, day 51) was not susceptible to amikacin (MIC, 32 mg/liter), and its MIC for colistin varied (0.5 to 8 mg/liter). Whole-genome sequencing of SMKP01 revealed that 17 of 20 antimicrobial resistance genes, including qnrB91 (a novel qnrB2 variant) and aac(6')-Ib-cr, were located on an 86.9-kb IncFII-IncQ plasmid. The qnrB91 conferred greater fluoroquinolone resistance than qnrB2 SMKP03 aac(6')-Ib-cr that possessed a gene mutation that resulted in an R102W substitution, namely, aac(6')-Ib-D179Y, made a greater contribution to amikacin resistance than did aac(6')-Ib-cr SMKP03 harbored a nonsense mutation in mutS, which encodes a DNA repair enzyme. Introduction of this mutation into SMKP01 (SMKP01mutS _A307T) resulted in a dramatic increase (>58-fold) in the frequency of spontaneous amikacin-resistant mutants relative to SMKP01, and the substantial mutants possessed aac(6')-Ib-D179Y SMKP01mutS _A307T exhibited an unstable MIC for colistin (0.5 to 8 mg/liter). The results demonstrate that a disruptive mutation in MutS, arising during the clinical course of an infection, created a platform for the acquisition of amikacin nonsusceptibility and colistin heteroresistance in multidrug-resistant K. pneumoniae, mediated by the elevated frequency of spontaneous mutations.IMPORTANCE The emergence of multidrug resistance in pathogens such as Klebsiella pneumoniae is of great clinical concern. Antimicrobial resistance sometimes arises during the course of an infection. Although many studies have reported the emergence of antimicrobial resistance and novel antimicrobial resistance genes in the clinical isolates, the identity of the bacterial factor(s) that generate this emergence is still unclear. We report that a disruptive mutation in MutS, arising during the clinical course of an infection, created a context for the acquisition of colistin resistance and the emergence of a novel variant of the amikacin resistance gene in multidrug-resistant K. pneumoniae via an increase in the frequency of spontaneous mutation. This observation is important for understanding how K. pneumoniae develops multidrug resistance during infection and could potentially lead to new antimicrobial treatments for high-risk pathological microbes.

Hybrid Inhibitors of DNA Gyrase A and B: Design, Synthesis and Evaluation

The discovery of multi-targeting ligands of bacterial enzymes is an important strategy to combat rapidly spreading antimicrobial resistance. Bacterial DNA gyrase and topoisomerase IV are validated targets for the development of antibiotics. They can be inhibited at their catalytic sites or at their ATP binding sites. Here we present the design of new hybrids between the catalytic inhibitor ciprofloxacin and ATP-competitive inhibitors that show low nanomolar inhibition of DNA gyrase and antibacterial activity against Gram-negative pathogens. The most potent hybrid 3a has MICs of 0.5 µg/mL against Klebsiella pneumoniae, 4 µg/mL against Enterobacter cloacae, and 2 µg/mL against Escherichia coli. In addition, inhibition of mutant E. coli strains shows that these hybrid inhibitors interact with both subunits of DNA gyrase (GyrA, GyrB), and that binding to both of these sites contributes to their antibacterial activity.

High-throughput laboratory evolution reveals evolutionary constraints in Escherichia coli

Understanding the constraints that shape the evolution of antibiotic resistance is critical for predicting and controlling drug resistance. Despite its importance, however, a systematic investigation of evolutionary constraints is lacking. Here, we perform a high-throughput laboratory evolution of Escherichia coli under the addition of 95 antibacterial chemicals and quantified the transcriptome, resistance, and genomic profiles for the evolved strains. Utilizing machine learning techniques, we analyze the phenotype–genotype data and identified low dimensional phenotypic states among the evolved strains. Further analysis reveals the underlying biological processes responsible for these distinct states, leading to the identification of trade-off relationships associated with drug resistance. We also report a decelerated evolution of β-lactam resistance, a phenomenon experienced by certain strains under various stresses resulting in higher acquired resistance to β-lactams compared to strains directly selected by β-lactams. These findings bridge the genotypic, gene expression, and drug resistance gap, while contributing to a better understanding of evolutionary constraints for antibiotic resistance.

Understanding evolutionary constraints in antibiotic resistance is crucial for prediction and control. Here, the authors use high-throughput laboratory evolution of Escherichia coli alongside machine learning to identify trade-off relationships associated with drug resistance.

Engineered biosynthetic pathways and biocatalytic cascades for sustainable synthesis

Nature exploits biosynthetic cascades to construct numerous molecules from a limited set of starting materials. A deeper understanding of biosynthesis and extraordinary developments in gene technology has allowed the manipulation of natural pathways and construction of artificial cascades for the preparation of a range of molecules, which would be challenging to access using traditional synthetic chemical approaches. Alongside these metabolic engineering strategies, there has been continued interest in developing in vivo and in vitro biocatalytic cascades. Advancements in both metabolic engineering and biocatalysis are complementary, and this article aims to highlight some of the most exciting developments in these two areas with a particular focus on exploring those that have the potential to advance both pathway engineering and more traditional biocatalytic cascade development.

Bacterial Genetic Engineering by Means of Recombineering for Reverse Genetics

Serving a robust platform for reverse genetics enabling the in vivo study of gene functions primarily in enterobacteriaceae, recombineering -or recombination-mediated genetic engineering-represents a powerful and relative straightforward genetic engineering tool. Catalyzed by components of bacteriophage-encoded homologous recombination systems and only requiring short ∼40–50 base homologies, the targeted and precise introduction of modifications (e.g., deletions, knockouts, insertions and point mutations) into the chromosome and other episomal replicons is empowered. Furthermore, by its ability to make use of both double- and single-stranded linear DNA editing substrates (e.g., PCR products or oligonucleotides, respectively), lengthy subcloning of specific DNA sequences is circumvented. Further, the more recent implementation of CRISPR-associated endonucleases has allowed for more efficient screening of successful recombinants by the selective purging of non-edited cells, as well as the creation of markerless and scarless mutants. In this review we discuss various recombineering strategies to promote different types of gene modifications, how they are best applied, and their possible pitfalls.

Underground isoleucine biosynthesis pathways in E. coli

The promiscuous activities of enzymes provide fertile ground for the evolution of new metabolic pathways. Here, we systematically explore the ability of E. coli to harness underground metabolism to compensate for the deletion of an essential biosynthetic pathway. By deleting all threonine deaminases, we generated a strain in which isoleucine biosynthesis was interrupted at the level of 2-ketobutyrate. Incubation of this strain under aerobic conditions resulted in the emergence of a novel 2-ketobutyrate biosynthesis pathway based upon the promiscuous cleavage of O-succinyl-L-homoserine by cystathionine γ-synthase (MetB). Under anaerobic conditions, pyruvate formate-lyase enabled 2-ketobutyrate biosynthesis from propionyl-CoA and formate. Surprisingly, we found this anaerobic route to provide a substantial fraction of isoleucine in a wild-type strain when propionate is available in the medium. This study demonstrates the selective advantage underground metabolism offers, providing metabolic redundancy and flexibility which allow for the best use of environmental carbon sources.

Unlocking nature's biosynthetic potential by directed genome evolution

Microorganisms have been increasingly explored as microbial cell factories for production of fuels, chemicals, drugs, and materials. Among the various metabolic engineering strategies, directed genome evolution has emerged as one of the most powerful tools to unlock the full biosynthetic potential of microorganisms. Here we summarize the directed genome evolution strategies that have been developed in recent years, including adaptive laboratory evolution and various targeted genome-scale engineering strategies, and discuss their applications in basic and applied biological research.

Release Factor Inhibiting Antimicrobial Peptides Improve Nonstandard Amino Acid Incorporation in Wild-type Bacterial Cells

We report a tunable chemical genetics approach for enhancing genetic code expansion in different wild-type bacterial strains that employ apidaecin-like, antimicrobial peptides observed to temporarily sequester and thereby inhibit Release Factor 1 (RF1). In a concentration-dependent matter, these peptides granted a conditional lambda phage resistance to a recoded Escherichia coli strain with nonessential RF1 activity and promoted multisite nonstandard amino acid (nsAA) incorporation at in-frame amber stop codons in vivo and in vitro. When exogenously added, the peptides stimulated specific nsAA incorporation in a variety of sensitive, wild-type (RF1+) strains, including Agrobacterium tumefaciens, a species in which nsAA incorporation has not been previously reported. Improvement in nsAA incorporation was typically 2-15-fold in E. coli BL21, MG1655, and DH10B strains and A. tumefaciens with the >20-fold improvement observed in probiotic E. coli Nissle 1917. In-cell expression of these peptides promoted multisite nsAA incorporation in transcripts with up to 6 amber codons, with a >35-fold increase in BL21 showing moderate toxicity. Leveraging this RF1 sensitivity allowed multiplexed partial recoding of MG1655 and DH10B that rapidly resulted in resistant strains that showed an additional approximately twofold boost to nsAA incorporation independent of the peptide. Finally, in-cell expression of an apidaecin-like peptide library allowed the discovery of new peptide variants with reduced toxicity that still improved multisite nsAA incorporation >25-fold. In parallel to genetic reprogramming efforts, these new approaches can facilitate genetic code expansion technologies in a variety of wild-type bacterial strains.

Targeted mutagenesis of multiple chromosomal regions in microbes

Directed evolution allows the effective engineering of proteins, biosynthetic pathways, and cellular functions. Traditional plasmid-based methods generally subject one or occasionally multiple genes-of-interest to mutagenesis, require time-consuming manual interventions, and the genes that are subjected to mutagenesis are outside of their native genomic context. Other methods mutagenize the whole genome unselectively which may distort the outcome. Recent recombineering- and CRISPR-based technologies radically change this field by allowing exceedingly high mutation rates at multiple, predefined loci in their native genomic context. In this review, we focus on recent technologies that potentially allow accelerated tunable mutagenesis at multiple genomic loci in the native genomic context of these target sequences. These technologies will be compared by four main criteria, including the scale of mutagenesis, portability to multiple microbial species, off-target mutagenesis, and cost-effectiveness. Finally, we discuss how these technical advances open new avenues in basic research and biotechnology.

Improved bacterial recombineering by parallelized protein discovery

Exploiting bacteriophage-derived homologous recombination processes has enabled precise, multiplex editing of microbial genomes and the construction of billions of customized genetic variants in a single day. The techniques that enable this, multiplex automated genome engineering (MAGE) and directed evolution with random genomic mutations (DIvERGE), are however, currently limited to a handful of microorganisms for which single-stranded DNA-annealing proteins (SSAPs) that promote efficient recombineering have been identified. Thus, to enable genome-scale engineering in new hosts, efficient SSAPs must first be found. Here we introduce a high-throughput method for SSAP discovery that we call "serial enrichment for efficient recombineering" (SEER). By performing SEER in Escherichia coli to screen hundreds of putative SSAPs, we identify highly active variants PapRecT and CspRecT. CspRecT increases the efficiency of single-locus editing to as high as 50% and improves multiplex editing by 5- to 10-fold in E. coli, while PapRecT enables efficient recombineering in Pseudomonas aeruginosa, a concerning human pathogen. CspRecT and PapRecT are also active in other, clinically and biotechnologically relevant enterobacteria. We envision that the deployment of SEER in new species will pave the way toward pooled interrogation of genotype-to-phenotype relationships in previously intractable bacteria.

I-Block: a simple Escherichia coli-based assay for studying sequence-specific DNA binding of proteins

We have developed a simple method called I-Block assay, which can detect sequence-specific binding of proteins to DNA in Escherichia coli. The method works by detecting competition between the protein of interest and RNA polymerase for binding to overlapping target sites in a plasmid-borne lacI promoter variant. The assay utilizes two plasmids and an E. coli host strain, from which the gene of the Lac repressor (lacI) has been deleted. One of the plasmids carries the lacI gene with a unique NheI restriction site created in the lacI promoter. The potential recognition sequences of the tested protein are inserted into the NheI site. Introduction of the plasmids into the E. coliΔlacI host represses the constitutive β-galactosidase synthesis of the host bacterium. If the studied protein expressed from a compatible plasmid binds to its target site in the lacI promoter, it will interfere with lacI transcription and lead to increased β-galactosidase activity. The method was tested with two zinc finger proteins, with the lambda phage cI857 repressor, and with CRISPR-dCas9 targeted to the lacI promoter. The I-Block assay was shown to work with standard liquid cultures, with cultures grown in microplate and with colonies on X-gal indicator plates.

Application of combinatorial optimization strategies in synthetic biology

In the first wave of synthetic biology, genetic elements, combined into simple circuits, are used to control individual cellular functions. In the second wave of synthetic biology, the simple circuits, combined into complex circuits, form systems-level functions. However, efforts to construct complex circuits are often impeded by our limited knowledge of the optimal combination of individual circuits. For example, a fundamental question in most metabolic engineering projects is the optimal level of enzymes for maximizing the output. To address this point, combinatorial optimization approaches have been established, allowing automatic optimization without prior knowledge of the best combination of expression levels of individual genes. This review focuses on current combinatorial optimization methods and emerging technologies facilitating their applications.

Rebooting Synthetic Phage-Inducible Chromosomal Islands: One Method to Forge Them All

Phage-inducible chromosomal islands (PICIs) are a widespread family of mobile genetic elements, which have an important role in bacterial pathogenesis. These elements mobilize among bacterial species at extremely high frequencies, representing an attractive tool for the delivery of synthetic genes. However, tools for their genetic manipulation are limited and timing consuming. Here, we have adapted a synthetic biology approach for rapidly editing of PICIs in Saccharomyces cerevisiae based on their ability to excise and integrate into the bacterial chromosome of their cognate host species. As proof of concept, we engineered several PICIs from Staphylococcus aureus and Escherichia coli and validated this methodology for the study of the biology of these elements by generating multiple and simultaneous mutations in different PICI genes. For biotechnological purposes, we also synthetically constructed PICIs as Trojan horses to deliver different CRISPR-Cas9 systems designed to either cure plasmids or eliminate cells carrying the targeted genes. Our results demonstrate that the strategy developed here can be employed universally to study PICIs and enable new approaches for diagnosis and treatment of bacterial diseases.

Escape mutations circumvent a tradeoff between resistance to a beta-lactam and resistance to a beta-lactamase inhibitor

Beta-lactamase inhibitors are increasingly used to counteract antibiotic resistance mediated by beta-lactamase enzymes. These inhibitors compete with the beta-lactam antibiotic for the same binding site on the beta-lactamase, thus generating an evolutionary tradeoff: mutations that increase the enzyme's beta-lactamase activity tend to increase also its susceptibility to the inhibitor. Here, we investigate how common and accessible are mutants that escape this adaptive tradeoff. Screening a deep mutant library of the bla_ampC beta-lactamase gene of Escherichia coli, we identified mutations that allow growth at beta-lactam concentrations far exceeding those inhibiting growth of the wildtype strain, even in the presence of the enzyme inhibitor (avibactam). These escape mutations are rare and drug-specific, and some combinations of avibactam with beta-lactam drugs appear to prevent such escape phenotypes. Our results, showing differential adaptive potential of bla_ampC to combinations of avibactam and different beta-lactam antibiotics, suggest that it may be possible to identify treatments that are more resilient to evolution of resistance.

Integrating CRISPR-Enabled Trackable Genome Engineering and Transcriptomic Analysis of Global Regulators for Antibiotic Resistance Selection and Identification in Escherichia coli

The growing threat of antimicrobial resistance poses a serious threat to public health care and motivates efforts to understand the means by which resistance acquisition occurs and how this can be combatted. To address these challenges, we expedited the identification of novel mutations that enable complex phenotypic changes that result in improved tolerance to antibiotics by integrating CREATE and transcriptomic analysis of global regulators. The results give us a better understanding of the mechanisms of resistance to tetracycline antibiotics and aminoglycoside antibiotics and also indicate that the method may be used for quickly identifying resistance-related mutations.

It is important to expedite our understanding of antibiotic resistance to address the increasing numbers of fatalities and environmental pollution due to the emergence of antibiotic resistance and multidrug-resistant strains. Here, we combined the CRISPR-enabled trackable genome engineering (CREATE) technology and transcriptomic analysis to investigate antibiotic tolerance in Escherichia coli. We developed rationally designed site saturation mutagenesis libraries targeting 23 global regulators to identify fitness-conferring mutations in response to diverse antibiotic stresses. We identified seven novel mutations that confer resistance to the ribosome-targeting antibiotics doxycycline, thiamphenicol, and gentamicin in E. coli. To the best of our knowledge, these mutations that we identified have not been reported previously during treatment with the indicated antibiotics. Transcriptome sequencing-based transcriptome analysis was further employed to evaluate the genome-wide changes in gene expression in E. coli for SoxR G121P and cAMP receptor protein (CRP) V140W reconstructions, and improved fitness in response to doxycycline and gentamicin was seen. In the case of doxycycline, we speculated that SoxR G121P significantly increased the expression of genes involved in carbohydrate metabolism and energy metabolism to promote cell growth for improved adaptation. In the CRP V140W mutant with improved gentamicin tolerance, the expression of several amino acid biosynthesis genes and fatty acid degradation genes was significantly changed, and these changes probably altered the cellular energy state to improve adaptation. These findings have important significance for understanding such nonspecific mechanisms of antibiotic resistance and developing new antibacterial drugs.

IMPORTANCE The growing threat of antimicrobial resistance poses a serious threat to public health care and motivates efforts to understand the means by which resistance acquisition occurs and how this can be combatted. To address these challenges, we expedited the identification of novel mutations that enable complex phenotypic changes that result in improved tolerance to antibiotics by integrating CREATE and transcriptomic analysis of global regulators. The results give us a better understanding of the mechanisms of resistance to tetracycline antibiotics and aminoglycoside antibiotics and also indicate that the method may be used for quickly identifying resistance-related mutations.

High-Efficiency Multi-site Genomic Editing of Pseudomonas putida through Thermoinducible ssDNA Recombineering

Application of single-stranded DNA recombineering for genome editing of species other than enterobacteria is limited by the efficiency of the recombinase and the action of endogenous mismatch repair (MMR) systems. In this work we have set up a genetic system for entering multiple changes in the chromosome of the biotechnologically relevant strain EM42 of Pseudomononas putida. To this end high-level heat-inducible co-transcription of the rec2 recombinase and P. putida's allele mutLE36KPP was designed under the control of the PL/cI857 system. Cycles of short thermal shifts followed by transformation with a suite of mutagenic oligos delivered different types of genomic changes at frequencies up to 10% per single modification. The same approach was instrumental to super-diversify short chromosomal portions for creating libraries of functional genomic segments-e.g., ribosomal-binding sites. These results enabled multiplexing of genome engineering of P. putida, as required for metabolic reprogramming of this important synthetic biology chassis.

Synthetic sequence entanglement augments stability and containment of genetic information in cells

In synthetic biology, methods for stabilizing genetically engineered functions and confining recombinant DNA to intended hosts are necessary to cope with natural mutation accumulation and pervasive lateral gene flow. We present a generalizable strategy to preserve and constrain genetic information through the computational design of overlapping genes. Overlapping a sequence with an essential gene altered its fitness landscape and produced a constrained evolutionary path, even for synonymous mutations. Embedding a toxin gene in a gene of interest restricted its horizontal propagation. We further demonstrated a multiplex and scalable approach to build and test >7500 overlapping sequence designs, yielding functional yet highly divergent variants from natural homologs. This work enables deeper exploration of natural and engineered overlapping genes and facilitates enhanced genetic stability and biocontainment in emerging applications.

Low mutational load and high mutation rate variation in gut commensal bacteria

Bacteria generally live in species-rich communities, such as the gut microbiota. Yet little is known about bacterial evolution in natural ecosystems. Here, we followed the long-term evolution of commensal Escherichia coli in the mouse gut. We observe the emergence of mutation rate polymorphism, ranging from wild-type levels to 1,000-fold higher. By combining experiments, whole-genome sequencing, and in silico simulations, we identify the molecular causes and explore the evolutionary conditions allowing these hypermutators to emerge and coexist within the microbiota. The hypermutator phenotype is caused by mutations in DNA polymerase III proofreading and catalytic subunits, which increase mutation rate by approximately 1,000-fold and stabilise hypermutator fitness, respectively. Strong mutation rate variation persists for >1,000 generations, with coexistence between lineages carrying 4 to >600 mutations. The in vivo molecular evolution pattern is consistent with fitness effects of deleterious mutations s_d ≤ 10⁻⁴/generation, assuming a constant effect or exponentially distributed effects with a constant mean. Such effects are lower than typical in vitro estimates, leading to a low mutational load, an inference that is observed in in vivo and in vitro competitions. Despite large numbers of deleterious mutations, we identify multiple beneficial mutations that do not reach fixation over long periods of time. This indicates that the dynamics of beneficial mutations are not shaped by constant positive Darwinian selection but could be explained by other evolutionary mechanisms that maintain genetic diversity. Thus, microbial evolution in the gut is likely characterised by partial sweeps of beneficial mutations combined with hitchhiking of slightly deleterious mutations, which take a long time to be purged because they impose a low mutational load. The combination of these two processes could allow for the long-term maintenance of intraspecies genetic diversity, including mutation rate polymorphism. These results are consistent with the pattern of genetic polymorphism that is emerging from metagenomics studies of the human gut microbiota, suggesting that we have identified key evolutionary processes shaping the genetic composition of this community.

Weak-effect deleterious mutations and negative frequency–dependent selection, acting on beneficial mutations, shape the dynamics of molecular evolution within the mouse gut microbiota.

Suppression of antibiotic resistance evolution by single-gene deletion

Antibiotic treatment generally results in the selection of resistant bacterial strains, and the dynamics of resistance evolution is dependent on complex interactions between cellular components. To better characterize the mechanisms of antibiotic resistance and evaluate its dependence on gene regulatory networks, we performed systematic laboratory evolution of Escherichia coli strains with single-gene deletions of 173 transcription factors under three different antibiotics. This resulted in the identification of several genes whose deletion significantly suppressed resistance evolution, including arcA and gutM. Analysis of double-gene deletion strains suggested that the suppression of resistance evolution caused by arcA and gutM deletion was not caused by epistatic interactions with mutations known to confer drug resistance. These results provide a methodological basis for combinatorial drug treatments that may help to suppress the emergence of resistant pathogens by inhibiting resistance evolution.

CRISPR/Cas9 recombineering-mediated deep mutational scanning of essential genes in Escherichia coli

Deep mutational scanning can provide significant insights into the function of essential genes in bacteria. Here, we developed a high-throughput method for mutating essential genes of Escherichia coli in their native genetic context. We used Cas9-mediated recombineering to introduce a library of mutations, created by error-prone PCR, within a gene fragment on the genome using a single gRNA pre-validated for high efficiency. Tracking mutation frequency through deep sequencing revealed biases in the position and the number of the introduced mutations. We overcame these biases by increasing the homology arm length and blocking mismatch repair to achieve a mutation efficiency of 85% for non-essential genes and 55% for essential genes. These experiments also improved our understanding of poorly characterized recombineering process using dsDNA donors with single nucleotide changes. Finally, we applied our technology to target rpoB, the beta subunit of RNA polymerase, to study resistance against rifampicin. In a single experiment, we validate multiple biochemical and clinical observations made in the previous decades and provide insights into resistance compensation with the study of double mutants.

Second-generation 4,5,6,7-tetrahydrobenzo[d]thiazoles as novel DNA gyrase inhibitors

Aim: DNA gyrase and topoisomerase IV are essential bacterial enzymes, and in the fight against bacterial resistance, they are important targets for the development of novel antibacterial drugs. Results: Building from our first generation of 4,5,6,7-tetrahydrobenzo[d]thiazole-based DNA gyrase inhibitors, we designed and prepared an optimized series of analogs that show improved inhibition of DNA gyrase and topoisomerase IV from Staphylococcus aureus and Escherichia coli, with IC₅₀ values in the nanomolar range. Importantly, these inhibitors also show improved antibacterial activity against Gram-positive strains. Conclusion: The most promising inhibitor, 29, is active against Enterococcus faecalis, Enterococcus faecium and S. aureus wild-type and resistant strains, with minimum inhibitory concentrations between 4 and 8 μg/ml, which represents good starting point for development of novel antibacterials.

Deciphering the role of the capsule of Klebsiella pneumoniae during pathogenesis: A cautionary tale

Extracellular capsule polysaccharides increase the cellular fitness under abiotic stresses and during competition with other bacteria. They are best-known for their role in virulence, particularly in human hosts. Specifically, capsules facilitate tissue invasion by enhancing bacterial evasion from phagocytosis and protect cells from biocidal molecules. Klebsiella pneumoniae is a worrisome nosocomial pathogen with few known virulence factors, but the most important one is its capsule. In this issue, Tan et al. assess the fitness advantage of the capsule by competing a wild-type strain against four different mutants where capsule production is interrupted at different stages of the biosynthetic pathway. Strikingly, not all mutants provide a fitness advantage. They suggest that some mutants have secondary defects altering virulence-associated phenotypes and blurring the role of the capsule in pathogenesis. This study indicates that the K1 capsule in K. pneumoniae is not required for gut colonization but that it is critical for bloodstream dissemination to other organs. These results contribute to clarify the contradictory literature on the role of the Klebsiella capsule during infection. Finally, the varying fitness effects of different capsule mutations observed for K. pneumoniae K1 might apply also to other capsulated diderm bacteria that are facultative or emerging pathogens.

Growth of E. coli on formate and methanol via the reductive glycine pathway

Engineering a biotechnological microorganism for growth on one-carbon intermediates, produced from the abiotic activation of CO₂, is a key synthetic biology step towards the valorization of this greenhouse gas to commodity chemicals. Here we redesign the central carbon metabolism of the model bacterium Escherichia coli for growth on one-carbon compounds using the reductive glycine pathway. Sequential genomic introduction of the four metabolic modules of the synthetic pathway resulted in a strain capable of growth on formate and CO₂ with a doubling time of ~70 h and growth yield of ~1.5 g cell dry weight (gCDW) per mol-formate. Short-term evolution decreased doubling time to less than 8 h and improved biomass yield to 2.3 gCDW per mol-formate. Growth on methanol and CO₂ was achieved by further expression of a methanol dehydrogenase. Establishing synthetic formatotrophy and methylotrophy, as demonstrated here, paves the way for sustainable bioproduction rooted in CO₂ and renewable energy.