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

Enhanced eMAGE applied to identify genetic factors of nuclear hormone receptor dysfunction via combinatorial gene editing

Technologies that generate precise combinatorial genome modifications are well suited to dissect the polygenic basis of complex phenotypes and engineer synthetic genomes. Genome modifications with engineered nucleases can lead to undesirable repair outcomes through imprecise homology-directed repair, requiring non-cleavable gene editing strategies. Eukaryotic multiplex genome engineering (eMAGE) generates precise combinatorial genome modifications in Saccharomyces cerevisiae without generating DNA breaks or using engineered nucleases. Here, we systematically optimize eMAGE to achieve 90% editing frequency, reduce workflow time, and extend editing distance to 20 kb. We further engineer an inducible dominant negative mismatch repair system, allowing for high-efficiency editing via eMAGE while suppressing the elevated background mutation rate 17-fold resulting from mismatch repair inactivation. We apply these advances to construct a library of cancer-associated mutations in the ligand-binding domains of human estrogen receptor alpha and progesterone receptor to understand their impact on ligand-independent autoactivation. We validate that this yeast model captures autoactivation mutations characterized in human breast cancer models and further leads to the discovery of several previously uncharacterized autoactivating mutations. This work demonstrates the development and optimization of a cleavage-free method of genome editing well suited for applications requiring efficient multiplex editing with minimal background mutations.

Synthetic genomes unveil the effects of synonymous recoding

Engineering the genetic code of an organism provides the basis for (i) making any organism safely resistant to natural viruses and (ii) preventing genetic information flow into and out of genetically modified organisms while (iii) allowing the biosynthesis of genetically encoded unnatural polymers1-4. Achieving these three goals requires the reassignment of multiple of the 64 codons nature uses to encode proteins. However, synonymous codon replacement-recoding-is frequently lethal, and how recoding impacts fitness remains poorly explored. Here, we explore these effects using whole-genome synthesis, multiplexed directed evolution, and genome-transcriptome-translatome-proteome co-profiling on multiple recoded genomes. Using this information, we assemble a synthetic Escherichia coli genome in seven sections using only 57 codons to encode proteins. By discovering the rules responsible for the lethality of synonymous recoding and developing a data-driven multi-omics-based genome construction workflow that troubleshoots synthetic genomes, we overcome the lethal effects of 62,007 synonymous codon swaps and 11,108 additional genomic edits. We show that synonymous recoding induces transcriptional noise including new antisense RNAs, leading to drastic transcriptome and proteome perturbation. As the elimination of select codons from an organism's genetic code results in the widespread appearance of cryptic promoters, we show that synonymous codon choice may naturally evolve to minimize transcriptional noise. Our work provides the first genome-scale description of how synonymous codon changes influence organismal fitness and paves the way for the construction of functional genomes that provide genetic firewalls from natural ecosystems and safely produce biopolymers, drugs, and enzymes with an expanded chemistry.

ORBIT for E. coli: kilobase-scale oligonucleotide recombineering at high throughput and high efficiency

Microbiology and synthetic biology depend on reverse genetic approaches to manipulate bacterial genomes; however, existing methods require molecular biology to generate genomic homology, suffer from low efficiency, and are not easily scaled to high throughput. To overcome these limitations, we developed a system for creating kilobase-scale genomic modifications that uses DNA oligonucleotides to direct the integration of a non-replicating plasmid. This method, Oligonucleotide Recombineering followed by Bxb-1 Integrase Targeting (ORBIT) was pioneered in Mycobacteria, and here we adapt and expand it for Escherichia coli. Our redesigned plasmid toolkit for oligonucleotide recombineering achieved significantly higher efficiency than λ Red double-stranded DNA recombineering and enabled precise, stable knockouts (≤134 kb) and integrations (≤11 kb) of various sizes. Additionally, we constructed multi-mutants in a single transformation, using orthogonal attachment sites. At high throughput, we used pools of targeting oligonucleotides to knock out nearly all known transcription factor and small RNA genes, yielding accurate, genome-wide, single mutant libraries. By counting genomic barcodes, we also show ORBIT libraries can scale to thousands of unique members (>30k). This work demonstrates that ORBIT for E. coli is a flexible reverse genetic system that facilitates rapid construction of complex strains and readily scales to create sophisticated mutant libraries.

Genome engineering of the human gut microbiome

The human gut microbiome, a complex ecosystem, significantly influences host health, impacting crucial aspects such as metabolism and immunity. To enhance our comprehension and control of the molecular mechanisms orchestrating the intricate interplay between gut commensal bacteria and human health, the exploration of genome engineering for gut microbes is a promising frontier. Nevertheless, the complexities and diversities inherent in the gut microbiome pose substantial challenges to the development of effective genome engineering tools for human gut microbes. In this comprehensive review, we provide an overview of the current progress and challenges in genome engineering of human gut commensal bacteria, whether executed in vitro or in situ. A specific focus is directed towards the advancements and prospects in cargo DNA delivery and high-throughput techniques. Additionally, we elucidate the immense potential of genome engineering methods to enhance our understanding of the human gut microbiome and engineer the microorganisms to enhance human health.

Automated in vivo enzyme engineering accelerates biocatalyst optimization

Achieving cost-competitive bio-based processes requires development of stable and selective biocatalysts. Their realization through in vitro enzyme characterization and engineering is mostly low throughput and labor-intensive. Therefore, strategies for increasing throughput while diminishing manual labor are gaining momentum, such as in vivo screening and evolution campaigns. Computational tools like machine learning further support enzyme engineering efforts by widening the explorable design space. Here, we propose an integrated solution to enzyme engineering challenges whereby ML-guided, automated workflows (including library generation, implementation of hypermutation systems, adapted laboratory evolution, and in vivo growth-coupled selection) could be realized to accelerate pipelines towards superior biocatalysts.

Proteins à la carte: riboproteogenomic exploration of bacterial N-terminal proteoform expression

In recent years, it has become evident that the true complexity of bacterial proteomes remains underestimated. Gene annotation tools are known to propagate biases and overlook certain classes of truly expressed proteins, particularly proteoforms-protein isoforms arising from a single gene. Recent (re-)annotation efforts heavily rely on ribosome profiling by providing a direct readout of translation to fully describe bacterial proteomes. In this study, we employ a robust riboproteogenomic pipeline to conduct a systematic census of expressed N-terminal proteoform pairs, representing two isoforms encoded by a single gene raised by annotated and alternative translation initiation, in Salmonella. Intriguingly, conditional-dependent changes in relative utilization of annotated and alternative translation initiation sites (TIS) were observed in several cases. This suggests that TIS selection is subject to regulatory control, adding yet another layer of complexity to our understanding of bacterial proteomes.

IMPORTANCE

With the emerging theme of genes within genes comprising the existence of alternative open reading frames (ORFs) generated by translation initiation at in-frame start codons, mechanisms that control the relative utilization of annotated and alternative TIS need to be unraveled and our molecular understanding of resulting proteoforms broadened. Utilizing complementary ribosome profiling strategies to map ORF boundaries, we uncovered dual-encoding ORFs generated by in-frame TIS usage in Salmonella. Besides demonstrating that alternative TIS usage may generate proteoforms with different characteristics, such as differential localization and specialized function, quantitative aspects of conditional retapamulin-assisted ribosome profiling (Ribo-RET) translation initiation maps offer unprecedented insights into the relative utilization of annotated and alternative TIS, enabling the exploration of gene regulatory mechanisms that control TIS usage and, consequently, the translation of N-terminal proteoform pairs.

Employing synthetic biology to expand antibiotic discovery

Antimicrobial-resistant (AMR) bacterial pathogens are a continually growing threat as our methods for combating these infections continue to be overcome by the evolution of resistance mechanisms. Recent therapeutic methods have not staved off the concern of AMR infections, so continued research focuses on new ways of identifying small molecules to treat AMR pathogens. While chemical modification of existing antibiotics is possible, there has been rapid development of resistance by pathogens that were initially susceptible to these compounds. Synthetic biology is becoming a key strategy in trying to predict and induce novel, natural antibiotics. Advances in cloning and mutagenesis techniques applied through a synthetic biology lens can help characterize the native regulation of antibiotic biosynthetic gene clusters (BGCs) to identify potential modifications leading to more potent antibiotic activity. Additionally, many cryptic antibiotic BGCs are derived from non-ribosomal peptide synthase (NRPS) and polyketide synthase (PKS) biosynthetic pathways; complex, clustered genetic sequences that give rise to amino acid-derived natural products. Synthetic biology can be applied to modify and metabolically engineer these enzyme-based systems to promote rapid and sustainable production of natural products and their variants. This review will focus on recent advances related to synthetic biology as applied to genetic pathway characterization and identification of antibiotics from naturally occurring BGCs. Specifically, we will summarize recent efforts to characterize BGCs via general genomic mutagenesis, endogenous gene expression, and heterologous gene expression.

Mutagenesis techniques for evolutionary engineering of microbes - exploiting CRISPR-Cas, oligonucleotides, recombinases, and polymerases

The natural process of evolutionary adaptation is often exploited as a powerful tool to obtain microbes with desirable traits. For industrial microbes, evolutionary engineering is often used to generate variants that show increased yields or resistance to stressful industrial environments, thus obtaining superior microbial cell factories. However, even in large populations, the natural supply of beneficial mutations is typically low, which implies that obtaining improved microbes is often time-consuming and inefficient. To overcome this limitation, different techniques have been developed that boost mutation rates. While some of these methods simply increase the overall mutation rate across a genome, others use recent developments in DNA synthesis, synthetic biology, and CRISPR-Cas techniques to control the type and location of mutations. This review summarizes the most important recent developments and methods in the field of evolutionary engineering in model microorganisms. It discusses how both in vitro and in vivo approaches can increase the genetic diversity of the host, with a special emphasis on in vivo techniques for the optimization of metabolic pathways for precision fermentation.

Multicolor optogenetics for regulating flux ratio of three glycolytic pathways using EL222 and CcaSR in Escherichia coli

Optogenetics is an attractive synthetic biology tool for controlling the metabolic flux distribution. Here, we demonstrated optogenetic flux ratio control of glycolytic pathways consisting of the Embden-Meyerhof-Parnas (EMP), pentose phosphate (PP), and Entner-Doudoroff (ED) pathways by illuminating multicolor lights using blue light-responsive EL222 and green/red light-responsive CcaSR in Escherichia coli. EL222 forms a dimer and binds to a particular DNA sequence under blue light; therefore, target gene expression can be reduced or induced by inserting a recognition sequence into its promoter regions. First, a flux ratio between the PP and ED pathways was controlled by blue light using EL222. After blocking the EMP pathway, the EL222-recognition sequence was inserted between the -35 and -10 regions of gnd to repress the PP flux and was also inserted upstream of the -35 region of edd to induce ED flux. After adjusting light intensity, the PP:ED flux ratios were 60:39% and 29:70% under dark and blue light conditions, respectively. Finally, a CcaSR-based pgi expression system was implemented to control the flux ratio between the EMP and PP + ED pathways by illuminating green/red light. The EMP:PP:ED flux ratios were 80:9:11%, 14:35:51%, and 33:5:62% under green, red, and red and blue light, respectively.

A convergent evolutionary pathway attenuating cellulose production drives enhanced virulence of some bacteria

Bacteria adapt to selective pressure in their immediate environment in multiple ways. One mechanism involves the acquisition of independent mutations that disable or modify a key pathway, providing a signature of adaptation via convergent evolution. Extra-intestinal pathogenic Escherichia coli (ExPEC) belonging to sequence type 95 (ST95) represent a global clone frequently associated with severe human infections including acute pyelonephritis, sepsis, and neonatal meningitis. Here, we analysed a publicly available dataset of 613 ST95 genomes and identified a series of loss-of-function mutations that disrupt cellulose production or its modification in 55.3% of strains. We show the inability to produce cellulose significantly enhances ST95 invasive infection in a rat model of neonatal meningitis, leading to the disruption of intestinal barrier integrity in newborn pups and enhanced dissemination to the liver, spleen and brain. Consistent with these observations, disruption of cellulose production in ST95 augmented innate immune signalling and tissue neutrophil infiltration in a mouse model of urinary tract infection. Mutations that disrupt cellulose production were also identified in other virulent ExPEC STs, Shigella and Salmonella, suggesting a correlative association with many Enterobacteriaceae that cause severe human infection. Together, our findings provide an explanation for the emergence of hypervirulent Enterobacteriaceae clones.

Interaction with a phage gene underlie costs of a β-lactamase

The fitness cost of an antibiotic resistance gene (ARG) can differ across host strains, creating refuges that allow the maintenance of an ARG in the absence of direct selection for its resistance phenotype. Despite the importance of such ARG-host interactions for predicting ARG dynamics, the basis of ARG fitness costs and their variability between hosts are not well understood. We determined the genetic basis of a host-dependent cost of a β-lactamase, blaTEM-116*, that conferred a significant cost in one Escherichia coli strain but was close to neutral in 11 other Escherichia spp. strains. Selection of a blaTEM-116*-encoding plasmid in the strain in which it initially had a high cost resulted in rapid and parallel compensation for that cost through mutations in a P1-like phage gene, relAP1. When the wild-type relAP1 gene was added to a strain in which it was not present and in which blaTEM-116* was neutral, it caused the ARG to become costly. Thus, relAP1 is both necessary and sufficient to explain blaTEM-116* costs in at least some host backgrounds. To our knowledge, these findings represent the first demonstrated case of the cost of an ARG being influenced by a genetic interaction with a phage gene. The interaction between a phage gene and a plasmid-borne ARG highlights the complexity of selective forces determining the maintenance and spread of ARGs and, by extension, encoding phage and plasmids in natural bacterial communities.IMPORTANCEAntibiotic resistance genes (ARGs) play a major role in the increasing problem of antibiotic resistance in clinically relevant bacteria. Selection of these genes occurs in the presence of antibiotics, but their eventual success also depends on the sometimes substantial costs they impose on host bacteria in antibiotic-free environments. We evolved an ARG that confers resistance to penicillin-type antibiotics in one host in which it did confer a cost and in one host in which it did not. We found that costs were rapidly and consistently reduced through parallel genetic changes in a gene encoded by a phage that was infecting the costly host. The unmutated version of this gene was sufficient to cause the ARG to confer a cost in a host in which it was originally neutral, demonstrating an antagonism between the two genetic elements and underlining the range and complexity of pressures determining ARG dynamics in natural populations.

Ultrahigh-throughput screening-assisted in vivo directed evolution for enzyme engineering

BACKGROUND

Classical directed evolution is a powerful approach for engineering biomolecules with improved or novel functions. However, it traditionally relies on labour- and time-intensive iterative cycles, due in part to the need for multiple molecular biology steps, including DNA transformation, and limited screening throughput.

RESULTS

In this study, we present an ultrahigh throughput in vivo continuous directed evolution system with thermosensitive inducible tunability, which is based on error-prone DNA polymerase expression modulated by engineered thermal-responsive repressor cI857, and genomic MutS mutant with temperature-sensitive defect for fixation of mutations in Escherichia coli. We demonstrated the success of the in vivo evolution platform with β-lactamase as a model, with an approximately 600-fold increase in the targeted mutation rate. Furthermore, the platform was combined with ultrahigh-throughput screening methods and employed to evolve α-amylase and the resveratrol biosynthetic pathway. After iterative rounds of enrichment, a mutant with a 48.3% improvement in α-amylase activity was identified via microfluidic droplet screening. In addition, when coupled with an in vivo biosensor in the resveratrol biosynthetic pathway, a variant with 1.7-fold higher resveratrol production was selected by fluorescence-activated cell sorting.

CONCLUSIONS

In this study, thermal-responsive targeted mutagenesis coupled with ultrahigh-throughput screening was developed for the rapid evolution of enzymes and biosynthetic pathways.

Comparison of phage-derived recombinases for genetic manipulation of Pseudomonas species

IMPORTANCE

The Pseudomonas genus contains many members currently being investigated for applications in biodegradation, biopesticides, biocontrol, and synthetic biology. Though several strains have been identified with beneficial properties, chromosomal manipulations to further improve these strains for commercial applications have been limited due to the lack of efficient genetic tools that have been tested across this genus. Here, we test the recombineering efficiencies of five phage-derived recombinases across three biotechnologically relevant Pseudomonas strains: P. putida KT2440, P. protegens Pf-5, and P. protegens CHA0. These results demonstrate a method to generate targeted mutations quickly and efficiently across these strains, ideally introducing a method that can be implemented across the Pseudomonas genus and a strategy that may be applied to develop analogous systems in other nonmodel bacteria.

Metagenomics harvested genus-specific single-stranded DNA-annealing proteins improve and expand recombineering in Pseudomonas species

The widespread Pseudomonas genus comprises a collection of related species with remarkable abilities to degrade plastics and polluted wastes and to produce a broad set of valuable compounds, ranging from bulk chemicals to pharmaceuticals. Pseudomonas possess characteristics of tolerance and stress resistance making them valuable hosts for industrial and environmental biotechnology. However, efficient and high-throughput genetic engineering tools have limited metabolic engineering efforts and applications. To improve their genome editing capabilities, we first employed a computational biology workflow to generate a genus-specific library of potential single-stranded DNA-annealing proteins (SSAPs). Assessment of the library was performed in different Pseudomonas using a high-throughput pooled recombinase screen followed by Oxford Nanopore NGS analysis. Among different active variants with variable levels of allelic replacement frequency (ARF), efficient SSAPs were found and characterized for mediating recombineering in the four tested species. New variants yielded higher ARFs than existing ones in Pseudomonas putida and Pseudomonas aeruginosa, and expanded the field of recombineering in Pseudomonas taiwanensisand Pseudomonas fluorescens. These findings will enhance the mutagenesis capabilities of these members of the Pseudomonas genus, increasing the possibilities for biotransformation and enhancing their potential for synthetic biology applications.  .

Structural basis for subversion of host cell actin cytoskeleton during Salmonella infection

Secreted bacterial type III secretion system (T3SS) proteins are essential for successful infection by many human pathogens. Both T3SS translocator SipC and effector SipA are critical for Salmonella infection by subversion of the host cell cytoskeleton, but the precise molecular interplay between them remains unknown. Here, using cryo-electron microscopy, we show that SipA binds along the F-actin grooves with a unique binding pattern. SipA stabilizes F-actin through charged interface residues and appears to prevent inorganic phosphate release through closure of the "back door" of adenosine 5'-triphosphate pocket. We also show that SipC enhances the binding of SipA to F-actin, thus demonstrating that a sequential presence of T3SS proteins in host cells is associated with a sequence of infection events-starting with actin nucleation, filament growth, and stabilization. Together, our data explain the coordinated interplay of a precisely tuned and highly effective mechanism during Salmonella infection and provide a blueprint for interfering with Salmonella effectors acting on actin.

Strategies to identify and edit improvements in synthetic genome segments episomally

Genome engineering projects often utilize bacterial artificial chromosomes (BACs) to carry multi-kilobase DNA segments at low copy number. However, all stages of whole-genome engineering have the potential to impose mutations on the synthetic genome that can reduce or eliminate the fitness of the final strain. Here, we describe improvements to a multiplex automated genome engineering (MAGE) protocol to improve recombineering frequency and multiplexability. This protocol was applied to recoding an Escherichia coli strain to replace seven codons with synonymous alternatives genome wide. Ten 44 402-47 179 bp de novo synthesized DNA segments contained in a BAC from the recoded strain were unable to complement deletion of the corresponding 33-61 wild-type genes using a single antibiotic resistance marker. Next-generation sequencing (NGS) was used to identify 1-7 non-recoding mutations in essential genes per segment, and MAGE in turn proved a useful strategy to repair these mutations on the recoded segment contained in the BAC when both the recoded and wild-type copies of the mutated genes had to exist by necessity during the repair process. Finally, two web-based tools were used to predict the impact of a subset of non-recoding missense mutations on strain fitness using protein structure and function calls.

Continuous Multiplexed Phage Genome Editing Using Recombitrons

Bacteriophages, which naturally shape bacterial communities, can be co-opted as a biological technology to help eliminate pathogenic bacteria from our bodies and food supply1. Phage genome editing is a critical tool to engineer more effective phage technologies. However, editing phage genomes has traditionally been a low efficiency process that requires laborious screening, counter selection, or in vitro construction of modified genomes2. These requirements impose limitations on the type and throughput of phage modifications, which in turn limit our knowledge and potential for innovation. Here, we present a scalable approach for engineering phage genomes using recombitrons: modified bacterial retrons3 that generate recombineering donor DNA paired with single stranded binding and annealing proteins to integrate those donors into phage genomes. This system can efficiently create genome modifications in multiple phages without the need for counterselection. Moreover, the process is continuous, with edits accumulating in the phage genome the longer the phage is cultured with the host, and multiplexable, with different editing hosts contributing distinct mutations along the genome of a phage in a mixed culture. In lambda phage, as an example, recombitrons yield single-base substitutions at up to 99% efficiency and up to 5 distinct mutations installed on a single phage genome, all without counterselection and only a few hours of hands-on time.

A swapped genetic code prevents viral infections and gene transfer

Engineering the genetic code of an organism has been proposed to provide a firewall from natural ecosystems by preventing viral infections and gene transfer1-6. However, numerous viruses and mobile genetic elements encode parts of the translational apparatus7-9, potentially rendering a genetic-code-based firewall ineffective. Here we show that such mobile transfer RNAs (tRNAs) enable gene transfer and allow viral replication in Escherichia coli despite the genome-wide removal of 3 of the 64 codons and the previously essential cognate tRNA and release factor genes. We then establish a genetic firewall by discovering viral tRNAs that provide exceptionally efficient codon reassignment allowing us to develop cells bearing an amino acid-swapped genetic code that reassigns two of the six serine codons to leucine during translation. This amino acid-swapped genetic code renders cells resistant to viral infections by mistranslating viral proteomes and prevents the escape of synthetic genetic information by engineered reliance on serine codons to produce leucine-requiring proteins. As these cells may have a selective advantage over wild organisms due to virus resistance, we also repurpose a third codon to biocontain this virus-resistant host through dependence on an amino acid not found in nature10. Our results may provide the basis for a general strategy to make any organism safely resistant to all natural viruses and prevent genetic information flow into and out of genetically modified organisms.

High-throughput genetic engineering tools for regulating gene expression in a microbial cell factory

With the rapid advances in biotechnological tools and strategies, microbial cell factory-constructing strategies have been established for the production of value-added compounds. However, optimizing the tradeoff between the biomass, yield, and titer remains a challenge in microbial production. Gene regulation is necessary to optimize and control metabolic fluxes in microorganisms for high-production performance. Various high-throughput genetic engineering tools have been developed for achieving rational gene regulation and genetic perturbation, diversifying the cellular phenotype and enhancing bioproduction performance. In this paper, we review the current high-throughput genetic engineering tools for gene regulation. In particular, technological approaches used in a diverse range of genetic tools for constructing microbial cell factories are introduced, and representative applications of these tools are presented. Finally, the prospects for high-throughput genetic engineering tools for gene regulation are discussed.

Plasticity and Stereotypic Rewiring of the Transcriptome Upon Bacterial Evolution of Antibiotic Resistance

Bacterial evolution of antibiotic resistance frequently has deleterious side effects on microbial growth, virulence, and susceptibility to other antimicrobial agents. However, it is unclear how these trade-offs could be utilized for manipulating antibiotic resistance in the clinic, not least because the underlying molecular mechanisms are poorly understood. Using laboratory evolution, we demonstrate that clinically relevant resistance mutations in Escherichia coli constitutively rewire a large fraction of the transcriptome in a repeatable and stereotypic manner. Strikingly, lineages adapted to functionally distinct antibiotics and having no resistance mutations in common show a wide range of parallel gene expression changes that alter oxidative stress response, iron homeostasis, and the composition of the bacterial outer membrane and cell surface. These common physiological alterations are associated with changes in cell morphology and enhanced sensitivity to antimicrobial peptides. Finally, the constitutive transcriptomic changes induced by resistance mutations are largely distinct from those induced by antibiotic stresses in the wild type. This indicates a limited role for genetic assimilation of the induced antibiotic stress response during resistance evolution. Our work suggests that diverse resistance mutations converge on similar global transcriptomic states that shape genetic susceptibility to antimicrobial compounds.

Reporting off-target effects of recombinant engineering using the pORTMAGE system

pORTMAGE recombineering is a simple technique for incorporation of novel point mutations into bacterial genomes that eliminates off-target effects. Here we inserted point mutations into the cusS gene from Escherichia coli, then, using Illumina sequencing, report genetic variants in all mutant strains. Several off-site mutations were found at high frequency. Low frequency mutations also show high heterogeneity. This means that it is essential for studies to report all off-target effects and acknowledge the effect that this may have on resultant phenotypes.

Biosensor-assisted titratable CRISPRi high-throughput (BATCH) screening for over-production phenotypes

With rapid advances in the development of metabolic pathways and synthetic biology toolkits, a persisting challenge in microbial bioproduction is how to optimally rewire metabolic fluxes and accelerate the concomitant high-throughput phenotype screening. Here we developed a biosensor-assisted titratable CRISPRi high-throughput (BATCH) screening approach that combines a titratable mismatch CRISPR interference and a biosensor mediated screening for high-production phenotypes in Escherichia coli. We first developed a programmable mismatch CRISPRi that could afford multiple levels of interference efficacy with a one-pot sgRNA pool (a total of 16 variants for each target gene) harboring two consecutive random mismatches in the seed region of sgRNA spacers. The mismatch CRISPRi was demonstrated to enable almost a full range of gene knockdown when targeting different positions on genes. As a proof-of-principle demonstration of the BATCH screening system, we designed doubly mismatched sgRNA pools targeting 20 relevant genes in E. coli and optimized a PadR-based p-coumaric acid biosensor with broad dynamic range for the eGFP fluorescence guided high-production screening. Using sgRNA variants for the combinatorial knockdown of pfkA and ptsI, the p-coumaric acid titer was increased by 40.6% to o 1308.6 mg/l from glycerol in shake flasks. To further demonstrate the general applicability of the BATCH screening system, we recruited a HpdR-based butyrate biosensor that facilitated the screening of E. coli strains achieving 19.0% and 25.2% increase of butyrate titer in shake flasks with sgRNA variants targeting sucA and ldhA, respectively. This work reported the establishment of a plug-and-play approach that enables multilevel modulation of metabolic fluxes and high-throughput screening of high-production phenotypes.

An efficient, scarless, selection-free technology for phage engineering

Most recently developed phage engineering technologies are based on the CRISPR-Cas system. Here, we present a non-CRISPR-based method for genetically engineering the Escherichia coli phages T5, T7, P1, and λ by adapting the pORTMAGE technology, which was developed for engineering bacterial genomes. The technology comprises E. coli harbouring a plasmid encoding a potent recombinase and a gene transiently silencing a repair system. Oligonucleotides with the desired phage mutation are electroporated into E. coli followed by infection of the target bacteriophage. The high efficiency of this technology, which yields 1-14% of desired recombinants, allows low-throughput screening for the desired mutant. We have demonstrated the use of this technology for single-base substitutions, for deletions of 50-201 bases, for insertions of 20 bases, and for four different phages. The technology may also be readily modified for use across many additional bacterial and phage strains.[Figure: see text].

Synthetic bacteria for the detection and bioremediation of heavy metals

Toxic heavy metal accumulation is one of anthropogenic environmental pollutions, which poses risks to human health and ecological systems. Conventional heavy metal remediation approaches rely on expensive chemical and physical processes leading to the formation and release of other toxic waste products. Instead, microbial bioremediation has gained interest as a promising and cost-effective alternative to conventional methods, but the genetic complexity of microorganisms and the lack of appropriate genetic engineering technologies have impeded the development of bioremediating microorganisms. Recently, the emerging synthetic biology opened a new avenue for microbial bioremediation research and development by addressing the challenges and providing novel tools for constructing bacteria with enhanced capabilities: rapid detection and degradation of heavy metals while enhanced tolerance to toxic heavy metals. Moreover, synthetic biology also offers new technologies to meet biosafety regulations since genetically modified microorganisms may disrupt natural ecosystems. In this review, we introduce the use of microorganisms developed based on synthetic biology technologies for the detection and detoxification of heavy metals. Additionally, this review explores the technical strategies developed to overcome the biosafety requirements associated with the use of genetically modified microorganisms.

Construction and application of an efficient dual-base editing platform for Bacillus subtilis evolution employing programmable base conversion

The functionally evolved bacterial chassis is of great importance to manufacture a group of assorted high value-added chemicals, from small molecules to biologically active macromolecules. However, the current evolution frameworks are less efficienct in generating in vivo genomic diversification because of insufficient tunability, rendering limited evolution spacing for chassis. Here, an engineered genomic diversification platform (CRISPR-ABE8e-CDA-nCas9) leveraging a programmable dual-deaminases base editor was fabricated for rapidly evolving bacterial chassis. The dual-base editor was constructed by reprogramming the CRISPR array, nCas9, and cytidine and adenosine deaminase, enabling single or multiple base conversion at the genomic scale by simultaneous C-to-T and A-to-G conversion in vivo. Employing titration of the Cas-deaminase fusion protein, the platform enabled editing any pre-defined genomic loci with tunable conversion efficiency and editable window, generating a repertoire of mutants with highly diversified genomic sequences. Leveraging the genomic diversification platform, we successfully evolved the nisin-resistant capability of Bacillus subtilis through directed evolution of the subunit of lantibiotic ATP-binding cassette. Therefore, our work provides a portable and programmable genomic diversification platform, which is promising to expedite the fabrication of high-performance and robust bacterial chassis used in the development of biomanufacturing and biopharmaceuticals.

Recombineering in Non-Model Bacteria

The technology of recombineering, in vivo genetic engineering, was initially developed in Escherichia coli and uses bacteriophage-encoded homologous recombination proteins to efficiently recombine DNA at short homologies (35 to 50 nt). Because the technology is homology driven, genomic DNA can be modified precisely and independently of restriction site location. Recombineering uses linear DNA substrates that are introduced into the cell by electroporation; these can be PCR products, synthetic double-strand DNA (dsDNA), or single-strand DNA (ssDNA). Here we describe the applications, challenges, and factors affecting ssDNA and dsDNA recombineering in a variety of non-model bacteria, both Gram-negative and -positive, and recent breakthroughs in the field. We list different microbes in which the widely used phage λ Red and Rac RecET recombination systems have been used for in vivo genetic engineering. New homologous ssDNA and dsDNA recombineering systems isolated from non-model bacteria are also described. The Basic Protocol outlines a method for ssDNA recombineering in the non-model species of Shewanella. The Alternate Protocol describes the use of CRISPR/Cas as a counter-selection system in conjunction with recombineering to enhance recovery of recombinants. We provide additional background information, pertinent considerations for experimental design, and parameters critical for success. The design of ssDNA oligonucleotides (oligos) and various internet-based tools for oligo selection from genome sequences are also described, as is the use of oligo-mediated recombination. This simple form of genome editing uses only ssDNA oligo(s) and does not require an exogenous recombination system. The information presented here should help researchers identify a recombineering system suitable for their microbe(s) of interest. If no system has been characterized for a specific microbe, researchers can find guidance in developing a recombineering system from scratch. We provide a flowchart of decision-making paths for strategically applying annealase-dependent or oligo-mediated recombination in non-model and undomesticated bacteria. © 2022 Wiley Periodicals LLC. This article has been contributed to by U.S. Government employees and their work is in the public domain in the USA. Basic Protocol: ssDNA recombineering in Shewanella species Alternate Protocol: ssDNA recombineering coupled to CRISPR/Cas9 in Shewanella species.

Steady-State Operation of a Cell-Free Genetic Band-Detection Circuit

Pattern formation processes play a decisive role during embryogenesis and involve the precise spatial and temporal orchestration of intricate gene regulatory processes. Synthetic gene circuits modeled after their biological counterparts can be used to investigate such processes under well-controlled conditions and may, in the future, be utilized for autonomous position determination in synthetic biological materials. Here, we investigated a three-node feed-forward gene regulatory circuit in vitro that generates three distinct fluorescent outputs in response to varying concentrations of a single externally supplied morphogen. The circuit acts as a band detector for the morphogen concentration and, in a spatial context, could serve as a stripe-forming gene circuit. We simulated the behavior of the genetic circuit in the presence of a morphogen gradient using a system of ordinary differential equations and determined optimal parameters for stripe-pattern formation using an evolutionary algorithm. To analyze the subcircuits of the system, we conducted cell-free characterization experiments and finally tested the whole genetic circuit in nanoliter-scale microfluidic flow reactors that provided a continuous supply of cell extract and metabolites and allowed removal of degradation products. To make use of the widely employed promoters P_LlacO-1 and P_LtetO-1 in our design, we removed LacI from our bacterial cell extract by genome engineering Escherichia coli using a pORTMAGE workflow. Our results show that the band-detector works as designed when operated out of equilibrium within the flow reactors. On the other hand, subcircuits of the system and the whole circuit fail to generate the desired gene expression response when operated in a closed reactor. Our work thus underlines the importance of out-of-equilibrium operation of complex gene circuits, which cannot settle to a steady-state expression pattern within the finite lifetime of a cell-free expression system.

SURE editing: combining oligo-recombineering and programmable insertion/deletion of selection markers to efficiently edit the Mycoplasma pneumoniae genome

The development of advanced genetic tools is boosting microbial engineering which can potentially tackle wide-ranging challenges currently faced by our society. Here we present SURE editing, a multi-recombinase engineering rationale combining oligonucleotide recombineering with the selective capacity of antibiotic resistance via transient insertion of selector plasmids. We test this method in Mycoplasma pneumoniae, a bacterium with a very inefficient native recombination machinery. Using SURE editing, we can seamlessly generate, in a single step, a wide variety of genome modifications at high efficiencies, including the largest possible deletion of this genome (30 Kb) and the targeted complementation of essential genes in the deletion of a region of interest. Additional steps can be taken to remove the selector plasmid from the edited area, to obtain markerless or even scarless edits. Of note, SURE editing is compatible with different site-specific recombinases for mediating transient plasmid integration. This battery of selector plasmids can be used to select different edits, regardless of the target sequence, which significantly reduces the cloning load associated to genome engineering projects. Given the proven functionality in several microorganisms of the machinery behind the SURE editing logic, this method is likely to represent a valuable advance for the synthetic biology field.

Unlocking the bacterial domain for industrial biotechnology applications using universal parts and tools

Synthetic biology can play a major role in the development of sustainable industrial biotechnology processes. However, the development of economically viable production processes is currently hampered by the limited availability of host organisms that can be engineered for a specific production process. To date, standard hosts such as Escherichia coli and Saccharomyces cerevisiae are often used as starting points for process development since parts and tools allowing their engineering are readily available. However, their suboptimal metabolic background or impaired performance at industrial scale for a desired production process, can result in increased costs associated with process development and/or disappointing production titres. Building a universal and portable gene expression system allowing genetic engineering of hosts across the bacterial domain would unlock the bacterial domain for industrial biotechnology applications in a highly standardized manner and doing so, render industrial biotechnology processes more competitive compared to the current polluting chemical processes. This review gives an overview of a selection of bacterial hosts highly interesting for industrial biotechnology based on both their metabolic and process optimization properties. Moreover, the requirements and progress made so far to enable universal, standardized, and portable gene expression across the bacterial domain is discussed.

Pre-existing chromosomal polymorphisms in pathogenic E. coli potentiate the evolution of resistance to a last-resort antibiotic

Bacterial pathogens show high levels of chromosomal genetic diversity, but the influence of this diversity on the evolution of antibiotic resistance by plasmid acquisition remains unclear. Here, we address this problem in the context of colistin, a 'last line of defence' antibiotic. Using experimental evolution, we show that a plasmid carrying the MCR-1 colistin resistance gene dramatically increases the ability of Escherichia coli to evolve high-level colistin resistance by acquiring mutations in lpxC, an essential chromosomal gene involved in lipopolysaccharide biosynthesis. Crucially, lpxC mutations increase colistin resistance in the presence of the MCR-1 gene, but decrease the resistance of wild-type cells, revealing positive sign epistasis for antibiotic resistance between the chromosomal mutations and a mobile resistance gene. Analysis of public genomic datasets shows that lpxC polymorphisms are common in pathogenic E. coli, including those carrying MCR-1, highlighting the clinical relevance of this interaction. Importantly, lpxC diversity is high in pathogenic E. coli from regions with no history of MCR-1 acquisition, suggesting that pre-existing lpxC polymorphisms potentiated the evolution of high-level colistin resistance by MCR-1 acquisition. More broadly, these findings highlight the importance of standing genetic variation and plasmid/chromosomal interactions in the evolutionary dynamics of antibiotic resistance.

Single mutation makes Escherichia coli an insect mutualist

Microorganisms often live in symbiosis with their hosts, and some are considered mutualists, where all species involved benefit from the interaction. How free-living microorganisms have evolved to become mutualists is unclear. Here we report an experimental system in which non-symbiotic Escherichia coli evolves into an insect mutualist. The stinkbug Plautia stali is typically associated with its essential gut symbiont, Pantoea sp., which colonizes a specialized symbiotic organ. When sterilized newborn nymphs were infected with E. coli rather than Pantoea sp., only a few insects survived, in which E. coli exhibited specific localization to the symbiotic organ and vertical transmission to the offspring. Through transgenerational maintenance with P. stali, several hypermutating E. coli lines independently evolved to support the host's high adult emergence and improved body colour; these were called 'mutualistic' E. coli. These mutants exhibited slower bacterial growth, smaller size, loss of flagellar motility and lack of an extracellular matrix. Transcriptomic and genomic analyses of 'mutualistic' E. coli lines revealed independent mutations that disrupted the carbon catabolite repression global transcriptional regulator system. Each mutation reproduced the mutualistic phenotypes when introduced into wild-type E. coli, confirming that single carbon catabolite repression mutations can make E. coli an insect mutualist. These findings provide an experimental system for future work on host-microbe symbioses and may explain why microbial mutualisms are omnipresent in nature.

Evolutionary Instability of Collateral Susceptibility Networks in Ciprofloxacin-Resistant Clinical Escherichia coli Strains

Collateral sensitivity and resistance occur when resistance development toward one antimicrobial either potentiates or deteriorates the effect of others. Previous reports on collateral effects on susceptibility focus on newly acquired resistance determinants and propose that novel treatment guidelines informed by collateral networks may reduce the evolution, selection, and spread of antimicrobial resistance. In this study, we investigate the evolutionary stability of collateral networks in five ciprofloxacin-resistant, clinical Escherichia coli strains. After 300 generations of experimental evolution without antimicrobials, we show complete fitness restoration in four of five genetic backgrounds and demonstrate evolutionary instability in collateral networks of newly acquired resistance determinants. We show that compensatory mutations reducing efflux expression are the main drivers destabilizing initial collateral networks and identify rpoS as a putative target for compensatory evolution. Our results add another layer of complexity to future predictions and clinical application of collateral networks.

Ucl fimbriae regulation and glycan receptor specificity contribute to gut colonisation by extra-intestinal pathogenic Escherichia coli

Extra-intestinal pathogenic Escherichia coli (ExPEC) belong to a critical priority group of antibiotic resistant pathogens. ExPEC establish gut reservoirs that seed infection of the urinary tract and bloodstream, but the mechanisms of gut colonisation remain to be properly understood. Ucl fimbriae are attachment organelles that facilitate ExPEC adherence. Here, we investigated cellular receptors for Ucl fimbriae and Ucl expression to define molecular mechanisms of Ucl-mediated ExPEC colonisation of the gut. We demonstrate differential expression of Ucl fimbriae in ExPEC sequence types associated with disseminated infection. Genome editing of strains from two common sequence types, F11 (ST127) and UTI89 (ST95), identified a single nucleotide polymorphism in the ucl promoter that changes fimbriae expression via activation by the global stress-response regulator OxyR, leading to altered gut colonisation. Structure-function analysis of the Ucl fimbriae tip-adhesin (UclD) identified high-affinity glycan receptor targets, with highest affinity for sialyllacto-N-fucopentose VI, a structure likely to be expressed on the gut epithelium. Comparison of the UclD adhesin to the homologous UcaD tip-adhesin from Proteus mirabilis revealed that although they possess a similar tertiary structure, apart from lacto-N-fucopentose VI that bound to both adhesins at low-micromolar affinity, they recognize different fucose- and glucose-containing oligosaccharides. Competitive surface plasmon resonance analysis together with co-structural investigation of UcaD in complex with monosaccharides revealed a broad-specificity glycan binding pocket shared between UcaD and UclD that could accommodate these interactions. Overall, our study describes a mechanism of adaptation that augments establishment of an ExPEC gut reservoir to seed disseminated infections, providing a pathway for the development of targeted anti-adhesion therapeutics.

ExPEC infection of the urinary tract and bloodstream is frequently seeded from an intestinal reservoir, necessitating an understanding of the mechanisms that promote gut colonisation. Here we employed molecular and structural approaches to define the regulation and function of ExPEC Ucl fimbriae as a gut colonisation factor. We describe how mutations in the non-coding regulatory region of the ucl promoter cause increased Ucl fimbriae expression and promote enhanced gut colonisation via tuned induction by a global regulator that senses oxygen stress. We further define the glycan receptor targets of Ucl fimbriae and characterise the structural features of the Ucl adhesin that facilitate these interactions. These findings explain how ExPEC can adapt to survival in the gut to seed extra-intestinal infection.

Scarless Recombineering of Phage in Lysogenic State

We present a scarless recombineering-based method for introducing multiple point mutations into the genome of a temperate phage. The method uses the λ Red recombineering system to promote exogenous ssDNA oligos to anneal on the prophage lagging strand during host genome replication. DNA repair is suppressed by inducing the expression of a dominant-negative mutant protein of the methyl-directed mismatch repair system. Screening for recombinant cells without a selection marker is feasible due to its high recombination frequency, estimated as more than 40% after six cycles. The method enables scarless editing of the genome of a bacteriophage in 4-5 days.

Hyperaccurate Ribosomes for Improved Genetic Code Reprogramming

The reprogramming of the genetic code through the introduction of noncanonical amino acids (ncAAs) has enabled exciting advances in synthetic biology and peptide drug discovery. Ribosomes that function with high efficiency and fidelity are necessary for all of these efforts, but for challenging ncAAs, the competing processes of near-cognate readthrough and peptidyl-tRNA dropoff can be issues. Here we uncover the surprising extent of these competing pathways in the PURE translation system using mRNAs encoding peptides with affinity tags at the N- and C-termini. We also show that hyperaccurate or error restrictive ribosomes with mutations in ribosomal protein S12 lead to significant improvements in yield and fidelity in the context of both canonical AAs and a challenging α,α-disubstituted ncAA. Hyperaccurate ribosomes also improve yields for quadruplet codon readthrough for a tRNA containing an expanded anticodon stem-loop, although they are not able to eliminate triplet codon reading by this tRNA. The impressive improvements in fidelity and the simplicity of introducing this mutation alongside other efforts to engineer the translation apparatus make hyperaccurate ribosomes an important advance for synthetic biology.

Recent advances in high-throughput metabolic engineering: Generation of oligonucleotide-mediated genetic libraries

The preparation of genetic libraries is an essential step to evolve microorganisms and study genotype-phenotype relationships by high-throughput screening/selection. As the large-scale synthesis of oligonucleotides becomes easy, cheap, and high-throughput, numerous novel strategies have been developed in recent years to construct high-quality oligo-mediated libraries, leveraging state-of-art molecular biology tools for genome editing and gene regulation. This review presents an overview of recent advances in creating and characterizing in vitro and in vivo genetic libraries, based on CRISPR/Cas, regulatory RNAs, and recombineering, primarily for Escherichia coli and Saccharomyces cerevisiae. These libraries' applications in high-throughput metabolic engineering, strain evolution and protein engineering are also discussed.

Efficient and Iterative Retron-Mediated in vivo Recombineering in E. coli

Recombineering is an important tool in gene editing, enabling fast, precise and highly specific in vivo modification of microbial genomes. Oligonucleotide-mediated recombineering via the in vivo production of single-stranded DNA can overcome the limitations of traditional recombineering methods that rely on the exogenous delivery of editing templates. By modifying a previously reported plasmid-based system for fully in vivo single-stranded DNA recombineering, we demonstrate iterative editing of independent loci by utilizing a temperature-sensitive origin of replication for easy curing of the editing plasmid from recombinant cells. Optimization of the promoters driving the expression of the system’s functional components, combined with targeted counterselection against unedited cells with Cas9 nuclease, enabled editing efficiencies of 90–100%. The addition of a dominant-negative mutL allele to the system allowed single-nucleotide edits that were otherwise unachievable due to mismatch repair. Finally, we tested alternative recombinases and found that efficiency significantly increased for some targets. Requiring only a single cloning step for retargeting, our system provides an easy-to-use method for rapid, efficient construction of desired mutants.

Graphical Abstract

Cross-kingdom expression of synthetic genetic elements promotes discovery of metabolites in the human microbiome

Small molecules encoded by biosynthetic pathways mediate cross-species interactions and harbor untapped potential, which has provided valuable compounds for medicine and biotechnology. Since studying biosynthetic gene clusters in their native context is often difficult, alternative efforts rely on heterologous expression, which is limited by host-specific metabolic capacity and regulation. Here, we describe a computational-experimental technology to redesign genes and their regulatory regions with hybrid elements for cross-species expression in Gram-negative and -positive bacteria and eukaryotes, decoupling biosynthetic capacity from host-range constraints to activate silenced pathways. These synthetic genetic elements enabled the discovery of a class of microbiome-derived nucleotide metabolites-tyrocitabines-from Lactobacillus iners. Tyrocitabines feature a remarkable orthoester-phosphate, inhibit translational activity, and invoke unexpected biosynthetic machinery, including a class of "Amadori synthases" and "abortive" tRNA synthetases. Our approach establishes a general strategy for the redesign, expression, mobilization, and characterization of genetic elements in diverse organisms and communities.

Genome editing via non-viral delivery platforms: current progress in personalized cancer therapy

Cancer is a severe disease that substantially jeopardizes global health. Although considerable efforts have been made to discover effective anti-cancer therapeutics, the cancer incidence and mortality are still growing. The personalized anti-cancer therapies present themselves as a promising solution for the dilemma because they could precisely destroy or fix the cancer targets based on the comprehensive genomic analyses. In addition, genome editing is an ideal way to implement personalized anti-cancer therapy because it allows the direct modification of pro-tumor genes as well as the generation of personalized anti-tumor immune cells. Furthermore, non-viral delivery system could effectively transport genome editing tools (GETs) into the cell nucleus with an appreciable safety profile. In this manuscript, the important attributes and recent progress of GETs will be discussed. Besides, the laboratory and clinical investigations that seek for the possibility of combining non-viral delivery systems with GETs for the treatment of cancer will be assessed in the scope of personalized therapy.

Research progress of pathway and genome evolution in microbes

Microbes can produce valuable natural products widely applied in medicine, food and other important fields. Nevertheless, it is usually challenging to achieve ideal industrial yields due to low production rate and poor toxicity tolerance. Evolution is a constant mutation and adaptation process used to improve strain performance. Generally speaking, the synthesis of natural products in microbes is often intricate, involving multiple enzymes or multiple pathways. Individual evolution of a certain enzyme often fails to achieve the desired results, and may lead to new rate-limiting nodes that affect the growth of microbes. Therefore, it is inevitable to evolve the biosynthetic pathways or the whole genome. Here, we reviewed the pathway-level evolution including multi-enzyme evolution, regulatory elements engineering, and computer-aided engineering, as well as the genome-level evolution based on several tools, such as genome shuffling and CRISPR/Cas systems. Finally, we also discussed the major challenges faced by in vivo evolution strategies and proposed some potential solutions.

inPOSE: A Flexible Toolbox for Chromosomal Cloning and Amplification of Bacterial Transgenes

Cloning the genes and operons encoding heterologous functions in bacterial hosts is now almost exclusively carried out using plasmid vectors. This has multiple drawbacks, including the need for constant selection and variation in copy numbers. The chromosomal integration of transgenes has always offered a viable alternative; however, to date, it has been of limited use due to its tedious nature and often being limited to a single copy. We introduce here a strategy that uses bacterial insertion sequences, which are the simplest autonomous transposable elements to insert and amplify genetic cargo into a bacterial chromosome. Transgene insertion can take place either as transposition or homologous recombination, and copy number amplification is achieved using controlled copy-paste transposition. We display the successful use of IS1 and IS3 for this purpose in Escherichia coli cells using various selection markers. We demonstrate the insertion of selectable genes, an unselectable gene and a five-gene operon in up to two copies in a single step. We continue with the amplification of the inserted cassette to double-digit copy numbers within two rounds of transposase induction and selection. Finally, we analyze the stability of the cloned genetic constructs in the lack of selection and find it to be superior to all investigated plasmid-based systems. Due to the ubiquitous nature of transposable elements, we believe that with proper design, this strategy can be adapted to numerous other bacterial species.

Generating Single Nucleotide Point Mutations in E. coli with the No-SCAR System

Genetic manipulation of microbial genomes is highly relevant for studying biological systems and the development of biotechnologies. In E. coli, λ-Red recombineering is one of the most widely used gene-editing methods, enabling site-specific insertions, deletions, and point mutations of any genomic locus. The no-SCAR system combines λ-Red recombineering with CRISPR/Cas9 for programmable selection of recombinant cells. Recombineering results in the transient production of heteroduplex DNA, as only one strand of DNA is initially altered, leaving the mismatched bases susceptible to repair by the host methyl-directed mismatch repair (MMR) system and reduces the efficiency of generating single nucleotide point mutations. Here we describe a method, where expression of cas9 and the MMR-inhibiting mutL_E32K variant are independently controlled by anhydrotetracycline- and cumate-inducible promoters from the pCas9CyMutL plasmid. Thus, MMR is selectively inhibited until recombinant cells have undergone replication and the desired mutation is permanently incorporated. By transiently inhibiting MMR, the accumulation of off-target mutations typically associated with MMR-deficient cell types is minimized. Methods for designing the editing template and sgRNA, cloning of the sgRNA, induction of λ-Red and MutL_E32K, the transformation of editing oligo, and induction of Cas9 for mutant selection are detailed within.

Lambda Red Recombineering of Bacteriophage in the Lysogenic State

We present a recombineering-based method for editing the genome of a temperate phage. The method uses the lambda Red recombination system to edit the genome of a lysogenized host with a prophage compatible with bacteriophage lambda. Linear DNA is used as the recombination substrate and antibiotic resistance is used as the basis for selection of recombinants. The method enables the genetic manipulation of a prophage in 3-5 days.

Recombineering in Staphylococcus aureus

Recombineering has proven to be an extraordinarily powerful and versatile approach for the modification of bacterial genomes, but has historically not been possible in the important opportunistic pathogen Staphylococcus aureus. After evaluating the activity of various recombinases in S. aureus, we developed methods for recombineering in that organism using synthetic, single-stranded DNA oligonucleotides. This approach can be coupled to CRISPR/Cas9-mediated lethal counterselection in order to improve the efficiency with which recombinant S. aureus are recovered, which is especially useful in instances where mutants lack a selectable phenotype. These methods provide a rapid, scalable, precise, and inexpensive means to engineer point mutations, variable-length deletions, and short insertions into the S. aureus genome.

Precise genome editing across kingdoms of life using retron-derived DNA

Exogenous DNA can be a template to precisely edit a cell’s genome. However, the delivery of in vitro-produced DNA to target cells can be inefficient, and low abundance of template DNA may underlie the low rate of precise editing. One potential tool to produce template DNA inside cells is a retron, a bacterial retroelement involved in phage defense. However, little effort has been directed at optimizing retrons to produce designed sequences. Here, we identify modifications to the retron non-coding RNA that result in more abundant reverse transcribed DNA. By testing architectures of the retron operon that enable efficient reverse transcription, we find that gains in DNA production are portable from prokaryotic to eukaryotic cells and result in more efficient genome editing. Finally, we show that retron RT-DNA can be used to precisely edit cultured human cells. These experiments provide a general framework to produce DNA using retrons for genome modification.

Acceleration of target production in co-culture by enhancing intermediate consumption through adaptive laboratory evolution

Co-culture is a promising way to alleviate metabolic burden by dividing the metabolic pathways into several modules and sharing the conversion processes with multiple strains. Since an intermediate is passed from the donor to the recipient via the extracellular environment, it is inevitably diluted. Therefore, enhancing the intermediate consumption rate is important for increasing target productivity. In the present study, we demonstrated the enhancement of mevalonate consumption in Escherichia coli by adaptive laboratory evolution and applied the evolved strain to isoprenol production in an E. coli (upstream: glucose to mevalonate)-E. coli (downstream: mevalonate to isoprenol) co-culture. An engineered mevalonate auxotroph strain was repeatedly sub-cultured in a synthetic medium supplemented with mevalonate, where the mevalonate concentration was decreased stepwise from 100 to 20 µM. In five parallel evolution experiments, all growth rates gradually increased, resulting in five evolved strains. Whole-genome re-sequencing and reverse engineering identified three mutations involved in enhancing mevalonate consumption. After introducing nudF gene for producing isoprenol, the isoprenol-producing parental and evolved strains were respectively co-cultured with a mevalonate-producing strain. At an inoculation ratio of 1:3 (upstream:downstream), isoprenol production using the evolved strain was 3.3 times higher than that using the parental strain.

The chemotherapeutic drug methotrexate selects for antibiotic resistance

Background

Understanding drivers of antibiotic resistance evolution is fundamental for designing optimal treatment strategies and interventions to reduce the spread of antibiotic resistance. Various cytotoxic drugs used in cancer chemotherapy have antibacterial properties, but how bacterial populations are affected by these selective pressures is unknown. Here we test the hypothesis that the widely used cytotoxic drug methotrexate affects the evolution and selection of antibiotic resistance.

Methods

First, we determined methotrexate susceptibility (IC₉₀) and selective abilities in a collection of Escherichia coli and Klebsiella pneumoniae strains with and without pre-existing trimethoprim resistance determinants. We constructed fluorescently labelled pairs of E. coli MG1655 differing only in trimethoprim resistance determinants and determined the minimum selective concentrations of methotrexate using flow-cytometry. We further used an experimental evolution approach to investigate the effects of methotrexate on de novo trimethoprim resistance evolution.

Findings

We show that methotrexate can select for acquired trimethoprim resistance determinants located on the chromosome or a plasmid. Additionally, methotrexate co-selects for genetically linked resistance determinants when present together with trimethoprim resistance on a multi-drug resistance plasmid. These selective effects occur at concentrations 40- to >320-fold below the methotrexate minimal inhibitory concentration.

Interpretation

Our results strongly suggest a selective role of methotrexate for virtually any antibiotic resistance determinant when present together with trimethoprim resistance on a multi-drug resistance plasmid. The presented results may have significant implications for patient groups strongly depending on effective antibiotic treatment.

Funding

PJJ was supported by 10.13039/100007465UiT The Arctic University of Norway and the Northern Norway Regional Health Authority (SFP1292–16/HNF1586–21) and JPI-EC-AMR (Project 271,176/H10). DIA was supported by the 10.13039/501100004359Swedish Research Council (grant 2017–01,527). The publication charges for this article have been funded by a grant from the publication fund of UiT The Arctic University of Norway.

Evolutionary repair reveals an unexpected role of the tRNA modification m1G37 in aminoacylation

The tRNA modification m¹G37, introduced by the tRNA methyltransferase TrmD, is thought to be essential for growth in bacteria because it suppresses translational frameshift errors at proline codons. However, because bacteria can tolerate high levels of mistranslation, it is unclear why loss of m¹G37 is not tolerated. Here, we addressed this question through experimental evolution of trmD mutant strains of Escherichia coli. Surprisingly, trmD mutant strains were viable even if the m¹G37 modification was completely abolished, and showed rapid recovery of growth rate, mainly via duplication or mutation of the proline-tRNA ligase gene proS. Growth assays and in vitro aminoacylation assays showed that G37-unmodified tRNA^Pro is aminoacylated less efficiently than m¹G37-modified tRNA^Pro, and that growth of trmD mutant strains can be largely restored by single mutations in proS that restore aminoacylation of G37-unmodified tRNA^Pro. These results show that inefficient aminoacylation of tRNA^Pro is the main reason for growth defects observed in trmD mutant strains and that proS may act as a gatekeeper of translational accuracy, preventing the use of error-prone unmodified tRNA^Pro in translation. Our work shows the utility of experimental evolution for uncovering the hidden functions of essential genes and has implications for the development of antibiotics targeting TrmD.