High efficiency mutagenesis, repair, and engineering of chromosomal DNA using single-stranded oligonucleotides

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.

Mutational Analysis of Redβ Single Strand Annealing Protein: Roles of the 14 Lysine Residues in DNA Binding and Recombination In Vivo

Redβ is a 261 amino acid protein from bacteriophage λ that promotes a single-strand annealing (SSA) reaction for repair of double-stranded DNA (dsDNA) breaks. While there is currently no high-resolution structure available for Redβ, models of its DNA binding domain (residues 1-188) have been proposed based on homology with human Rad52, and a crystal structure of its C-terminal domain (CTD, residues 193-261), which binds to λ exonuclease and E. coli single-stranded DNA binding protein (SSB), has been determined. To evaluate these models, the 14 lysine residues of Redβ were mutated to alanine, and the variants tested for recombination in vivo and DNA binding and annealing in vitro. Most of the lysines within the DNA binding domain, including K36, K61, K111, K132, K148, K154, and K172, were found to be critical for DNA binding in vitro and recombination in vivo. By contrast, none of the lysines within the CTD, including K214, K245, K251, K253, and K258 were required for DNA binding in vitro, but two, K214 and K253, were critical for recombination in vivo, likely due to their involvement in binding to SSB. K61 was identified as a residue that is critical for DNA annealing, but not for initial ssDNA binding, suggesting a role in binding to the second strand of DNA incorporated into the complex. The K148A variant, which has previously been shown to be defective in oligomer formation, had the lowest affinity for ssDNA, and was the only variant that was completely non-cooperative, suggesting that ssDNA binding is coupled to oligomerization.

Resistance/fitness trade-off is a barrier to the evolution of MarR inactivation mutants in Escherichia coli

Background

Mutations that inactivate MarR reduce susceptibility to ciprofloxacin and competitive growth fitness in Escherichia coli. Both phenotypes are caused by overexpression of the MarA regulon, which includes the AcrAB-TolC drug efflux pump.

Objectives

We asked whether compensatory evolution could reduce the fitness cost of MarR-inactivating mutations without affecting resistance to ciprofloxacin.

Methods

The cost of overexpressing the AcrAB-TolC efflux pump was measured independently of MarA overexpression. Experimental evolution of MarR-inactive strains was used to select mutants with increased fitness. The acquired mutations were identified and their effects on drug susceptibility were measured.

Results

Overexpression of the AcrAB-TolC efflux pump was found not to contribute to the fitness cost of MarA regulon overexpression. Fitness-compensatory mutations were selected in marA and lon. The mutations reduced the level of MarA protein thus reducing expression of the MarA regulon. They restored growth fitness but also reduced resistance to ciprofloxacin.

Conclusions

The fitness cost caused by overexpression of the MarA regulon has multiple contributing factors. Experimental evolution did not identify any single pump-independent cost factor. Instead, efficient fitness compensation occurred only by mechanisms that reduce MarA concentration, which simultaneously reduce the drug resistance phenotype. This resistance/fitness trade-off is a barrier to the successful spread of MarR inactivation mutations in clinical isolates where growth fitness is essential.

Mismatch Intolerance of 5'-Truncated sgRNAs in CRISPR/Cas9 Enables Efficient Microbial Single-Base Genome Editing

The CRISPR/Cas9 system has recently emerged as a useful gene-specific editing tool. However, this approach occasionally results in the digestion of both the DNA target and similar DNA sequences due to mismatch tolerance, which remains a significant drawback of current genome editing technologies. However, our study determined that even single-base mismatches between the target DNA and 5'-truncated sgRNAs inhibited target recognition. These results suggest that a 5'-truncated sgRNA/Cas9 complex could be used to negatively select single-base-edited targets in microbial genomes. Moreover, we demonstrated that the 5'-truncated sgRNA method can be used for simple and effective single-base editing, as it enables the modification of individual bases in the DNA target, near and far from the 5' end of truncated sgRNAs. Further, 5'-truncated sgRNAs also allowed for efficient single-base editing when using an engineered Cas9 nuclease with an expanded protospacer adjacent motif (PAM; 5'-NG), which may enable whole-genome single-base editing.

Synthetic Genomes

DNA synthesis technology has progressed to the point that it is now practical to synthesize entire genomes. Quite a variety of methods have been developed, first to synthesize single genes but ultimately to massively edit or write from scratch entire genomes. Synthetic genomes can essentially be clones of native sequences, but this approach does not teach us much new biology. The ability to endow genomes with novel properties offers special promise for addressing questions not easily approachable with conventional gene-at-a-time methods. These include questions about evolution and about how genomes are fundamentally wired informationally, metabolically, and genetically. The techniques and technologies relating to how to design, build, and deliver big DNA at the genome scale are reviewed here. A fuller understanding of these principles may someday lead to the ability to truly design genomes from scratch.

Chemical-genetic interrogation of RNA polymerase mutants reveals structure-function relationships and physiological tradeoffs

The multi-subunit bacterial RNA polymerase (RNAP) and its associated regulators carry out transcription and integrate myriad regulatory signals. Numerous studies have interrogated RNAP mechanism, and RNAP mutations drive Escherichia coli adaptation to many health- and industry-relevant environments, yet a paucity of systematic analyses hampers our understanding of the fitness trade-offs from altering RNAP function. Here, we conduct a chemical-genetic analysis of a library of RNAP mutants. We discover phenotypes for non-essential insertions, show that clustering mutant phenotypes increases their predictive power for drawing functional inferences, and demonstrate that some RNA polymerase mutants both decrease average cell length and prevent killing by cell-wall targeting antibiotics. Our findings demonstrate that RNAP chemical-genetic interactions provide a general platform for interrogating structure-function relationships in vivo and for identifying physiological trade-offs of mutations, including those relevant for disease and biotechnology. This strategy should have broad utility for illuminating the role of other important protein complexes.

Phenotypic and genetic barriers to establishment of horizontally transferred genes encoding ribosomal protection proteins

Background

Ribosomal protection proteins (RPPs) interact with bacterial ribosomes to prevent inhibition of protein synthesis by tetracycline. RPP genes have evolved from a common ancestor into at least 12 distinct classes and spread by horizontal genetic transfer into a wide range of bacteria. Many bacterial genera host RPP genes from multiple classes but tet(M) is the predominant RPP gene found in Escherichia coli.

Objectives

We asked whether phenotypic barriers (low-level resistance, high fitness cost) might constrain the fixation of other RPP genes in E. coli.

Methods

We expressed a diverse set of six different RPP genes in E. coli, including tet(M), and quantified tetracycline susceptibility and growth phenotypes as a function of expression level, and evolvability to overcome identified phenotypic barriers.

Results

The genes tet(M) and tet(Q) conferred high-level tetracycline resistance without reducing fitness; tet(O) and tet(W) conferred high-level resistance but significantly reduced growth fitness; tetB(P) conferred low-level resistance and while mutants conferring high-level resistance were selectable these had reduced growth fitness; otr(A) did not confer resistance and resistant mutants could not be selected. Evolution experiments suggested that codon usage patterns in tet(O) and tet(W), and transcriptional silencing associated with nucleotide composition in tetB(P), accounted for the observed phenotypic barriers.

Conclusions

With the exception of tet(Q), the data reveal significant phenotypic and genetic barriers to the fixation of additional RPP genes in E. coli.

Expression of the qepA1 gene is induced under antibiotic exposure

Background

The qepA1 gene encodes an efflux pump that reduces susceptibility to ciprofloxacin. Little is known about the regulation of qepA1 expression.

Objectives

To assess the potential role of ciprofloxacin and other antibiotics in the regulation of qepA1 gene expression. To identify the promoter that drives qepA1 expression and other factors involved in expression regulation. To assess whether the identified features are universal among qepA alleles.

Methods

A translational qepA1-yfp fusion under the control of the qepA1 upstream region was cloned into the Escherichia coli chromosome. Expression of the fusion protein was measured in the presence of various antibiotics. Deletions within the upstream region were introduced to identify regions involved in gene expression and regulation. The qepA1 coding sequence and upstream region were compared with all available qepA sequences.

Results

Cellular stress caused by the presence of various antibiotics can induce qepA1 expression. The qepA1 gene is fused to a class I integron and gene expression is driven by the Pc promoter within the integrase gene. A segment within the integron belonging to a truncated dfrB4 gene is essential for the regulation of qepA1 expression. This genetic context is universal among all sequenced qepA alleles.

Conclusions

The fusion of the qepA1 gene to a class I integron has created a novel regulatory unit that enables qepA1 expression to be under the control of antibiotic exposure. This setup mitigates potential negative effects of QepA1 production on bacterial fitness by restricting high-level expression to environmental conditions in which QepA1 is beneficial.

Oligomeric complexes formed by Redβ single strand annealing protein in its different DNA bound states

Redβ is a single strand annealing protein from bacteriophage λ that binds loosely to ssDNA, not at all to pre-formed dsDNA, but tightly to a duplex intermediate of annealing. As viewed by electron microscopy, Redβ forms oligomeric rings on ssDNA substrate, and helical filaments on the annealed duplex intermediate. However, it is not clear if these are the functional forms of the protein in vivo. We have used size-exclusion chromatography coupled with multi-angle light scattering, analytical ultracentrifugation and native mass spectrometry (nMS) to characterize the size of the oligomers formed by Redβ in its different DNA-bound states. The nMS data, which resolve species with the highest resolution, reveal that Redβ forms an oligomer of 12 subunits in the absence of DNA, complexes ranging from 4 to 14 subunits on 38-mer ssDNA, and a much more distinct and stable complex of 11 subunits on 38-mer annealed duplex. We also measure the concentration of Redβ in cells active for recombination and find it to range from 7 to 27 μM. Collectively, these data provide new insights into the dynamic nature of the complex on ssDNA, and the more stable and defined complex on annealed duplex.

Rapid Genome Modification in Serratia marcescens Through Red Homologous Recombination

Despite the great potential of Serratia marcescens in industrial applications, lack of powerful genetic modification tools limits understanding of the regulatory networks of the useful metabolites and therefore restricts their mass production. To meet the urgent demand, we established a genome-editing strategy for S. marcescens based on Red recombineering in this study. Without host modification in advance, nucA and pigA were substituted by PCR-amplified resistance genes. No long homologous arms were required at the two sides of resistance genes. Using this procedure, the fragment at the S. marcescens as large as 20 kb was easily deleted. Then we constructed a counter-selection gene kil constructed under the control of inducible P_BAD operon, which demonstrates obvious lethality to S. marcescens. Subsequently, Gm^R-kil double selection cassette was inserted into the CDS of pigA gene. Using single-stranded DNA-mediated recombination, this insertion mutation was efficiently repaired through kil counter-selection. A powerful genetic modification platform based on Red recombineering system was successfully established for S. marcescens. Multiple types of modification and multiple recombination strategies can all be performed easily in this species. We hope this study will be useful for the theoretical research and the research of metabolic engineering in S. marcescens.

Engineered Bacteriophage Therapeutics: Rationale, Challenges and Future

The current problems with increasing bacterial resistance to antibacterial therapies, resulting in a growing frequency of incurable bacterial infections, necessitates the acceleration of studies on antibacterials of a new generation that could offer an alternative to antibiotics or support their action. Bacteriophages (phages) can kill antibiotic-sensitive as well as antibiotic-resistant bacteria, and thus are a major subject of such studies. Their efficacy in curing bacterial infections has been demonstrated in in vivo experiments and in the clinic. Unlike antibiotics, phages have a narrow range of specificity, which makes them safe for commensal microbiota. However, targeting even only the most clinically relevant strains of pathogenic bacteria requires large collections of well characterized phages, whose specificity would cover all such strains. The environment is a rich source of diverse phages, but due to their complex relationships with bacteria and safety concerns, only some naturally occurring phages can be considered for therapeutic applications. Still, their number and diversity make a detailed characterization of all potentially promising phages virtually impossible. Moreover, no single phage combines all the features required of an ideal therapeutic agent. Additionally, the rapid acquisition of phage resistance by bacteria may make phages already approved for therapy ineffective and turn the search for environmental phages of better efficacy and new specificity into an endless race. An alternative strategy for acquiring phages with desired properties in a short time with minimal cost regarding their acquisition, characterization, and approval for therapy could be based on targeted genome modifications of phage isolates with known properties. The first example demonstrating the potential of this strategy in curing bacterial diseases resistant to traditional therapy is the recent successful treatment of a progressing disseminated Mycobacterium abscessus infection in a teenage patient with the use of an engineered phage. In this review, we briefly present current methods of phage genetic engineering, highlighting their advantages and disadvantages, and provide examples of genetically engineered phages with a modified host range, improved safety or antibacterial activity, and proven therapeutic efficacy. We also summarize novel uses of engineered phages not only for killing pathogenic bacteria, but also for in situ modification of human microbiota to attenuate symptoms of certain bacterial diseases and metabolic, immune, or mental disorders.

Expanding Diversity of Firmicutes Single-Strand Annealing Proteins: A Putative Role of Bacteriophage-Host Arms Race

Bacteriophage-encoded single strand annealing proteins (SSAPs) are recombinases which can substitute the classical, bacterial RecA and manage the DNA metabolism at different steps of phage propagation. SSAPs have been shown to efficiently promote recombination between short and rather divergent DNA sequences and were exploited for in vivo genetic engineering mainly in Gram-negative bacteria. In opposition to the conserved and almost universal bacterial RecA protein, SSAPs display great sequence diversity. The importance for SSAPs in phage biology and phage-bacteria evolution is underlined by their role as key players in events of horizontal gene transfer (HGT). All of the above provoke a constant interest for the identification and study of new phage recombinase proteins in vivo, in vitro as well as in silico. Despite this, a huge body of putative ssap genes escapes conventional classification, as they are not properly annotated. In this work, we performed a wide-scale identification, classification and analysis of SSAPs encoded by the Firmicutes bacteria and their phages. By using sequence similarity network and gene context analyses, we created a new high quality dataset of phage-related SSAPs, substantially increasing the number of annotated SSAPs. We classified the identified SSAPs into seven distinct families, namely RecA, Gp2.5, RecT/Redβ, Erf, Rad52/22, Sak3, and Sak4, organized into three superfamilies. Analysis of the relationships between the revealed protein clusters led us to recognize Sak3-like proteins as a new distinct SSAP family. Our analysis showed an irregular phylogenetic distribution of ssap genes among different bacterial phyla and specific phages, which can be explained by the high rates of ssap HGT. We propose that the evolution of phage recombinases could be tightly linked to the dissemination of bacterial phage-resistance mechanisms (e.g., abortive infection and CRISPR/Cas systems) targeting ssap genes and be a part of the constant phage-bacteria arms race.

Genetic Architecture and Fitness of Bacterial Interspecies Hybrids

Integration of a conjugative plasmid into a bacterial chromosome can promote the transfer of chromosomal DNA to other bacteria. Intraspecies chromosomal conjugation is believed responsible for creating the global pathogens Klebsiella pneumoniae ST258 and Escherichia coli ST1193. Interspecies conjugation is also possible but little is known about the genetic architecture or fitness of such hybrids. To study this, we generated by conjugation 14 hybrids of E. coli and Salmonella enterica. These species belong to different genera, diverged from a common ancestor >100 Ma, and share a conserved order of orthologous genes with ∼15% nucleotide divergence. Genomic analysis revealed that all but one hybrid had acquired a contiguous segment of donor E. coli DNA, replacing a homologous region of recipient Salmonella chromosome, and ranging in size from ∼100 to >4,000 kb. Recombination joints occurred in sequences with higher-than-average nucleotide identity. Most hybrid strains suffered a large reduction in growth rate, but the magnitude of this cost did not correlate with the length of foreign DNA. Compensatory evolution to ameliorate the cost of low-fitness hybrids pointed towards disruption of complex genetic networks as a cause. Most interestingly, 4 of the 14 hybrids, in which from 45% to 90% of the Salmonella chromosome was replaced with E. coli DNA, showed no significant reduction in growth fitness. These data suggest that the barriers to creating high-fitness interspecies hybrids may be significantly lower than generally appreciated with implications for the creation of novel species.

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.

Comparing treatment strategies to reduce antibiotic resistance in an in vitro epidemiological setting

The rapid rise of antibiotic resistance, combined with the increasing cost and difficulties to develop new antibiotics, calls for treatment strategies that enable more sustainable antibiotic use. The development of such strategies, however, is impeded by the lack of suitable experimental approaches that allow testing their effects under realistic epidemiological conditions. Here, we present an approach to compare the effect of alternative multidrug treatment strategies in vitro using a robotic liquid-handling platform. We use this framework to study resistance evolution and spread implementing epidemiological population dynamics for treatment, transmission, and patient admission and discharge, as may be observed in hospitals. We perform massively parallel experimental evolution over up to 40 d and complement this with a computational model to infer the underlying population-dynamical parameters. We find that in our study, combination therapy outperforms monotherapies, as well as cycling and mixing, in minimizing resistance evolution and maximizing uninfecteds, as long as there is no influx of double resistance into the focal treated community.

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.

Approaches to genetic tool development for rapid domestication of non-model microorganisms

Non-model microorganisms often possess complex phenotypes that could be important for the future of biofuel and chemical production. They have received significant interest the last several years, but advancement is still slow due to the lack of a robust genetic toolbox in most organisms. Typically, "domestication" of a new non-model microorganism has been done on an ad hoc basis, and historically, it can take years to develop transformation and basic genetic tools. Here, we review the barriers and solutions to rapid development of genetic transformation tools in new hosts, with a major focus on Restriction-Modification systems, which are a well-known and significant barrier to efficient transformation. We further explore the tools and approaches used for efficient gene deletion, DNA insertion, and heterologous gene expression. Finally, more advanced and high-throughput tools are now being developed in diverse non-model microbes, paving the way for rapid and multiplexed genome engineering for biotechnology.

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.

Genome recoding strategies to improve cellular properties: mechanisms and advances

The genetic code, once believed to be universal and immutable, is now known to contain many variations and is not quite universal. The basis for genome recoding strategy is genetic code variation that can be harnessed to improve cellular properties. Thus, genome recoding is a promising strategy for the enhancement of genome flexibility, allowing for novel functions that are not commonly documented in the organism in its natural environment. Here, the basic concept of genetic code and associated mechanisms for the generation of genetic codon variants, including biased codon usage, codon reassignment, and ambiguous decoding, are extensively discussed. Knowledge of the concept of natural genetic code expansion is also detailed. The generation of recoded organisms and associated mechanisms with basic targeting components, including aminoacyl-tRNA synthetase-tRNA pairs, elongation factor EF-Tu and ribosomes, are highlighted for a comprehensive understanding of this concept. The research associated with the generation of diverse recoded organisms is also discussed. The success of genome recoding in diverse multicellular organisms offers a platform for expanding protein chemistry at the biochemical level with non-canonical amino acids, genetically isolating the synthetic organisms from the natural ones, and fighting viruses, including SARS-CoV2, through the creation of attenuated viruses. In conclusion, genome recoding can offer diverse applications for improving cellular properties in the genome-recoded organisms.

Reprogramming the genetic code

The encoded biosynthesis of proteins provides the ultimate paradigm for high-fidelity synthesis of long polymers of defined sequence and composition, but it is limited to polymerizing the canonical amino acids. Recent advances have built on genetic code expansion - which commonly permits the cellular incorporation of one type of non-canonical amino acid into a protein - to enable the encoded incorporation of several distinct non-canonical amino acids. Developments include strategies to read quadruplet codons, use non-natural DNA base pairs, synthesize completely recoded genomes and create orthogonal translational components with reprogrammed specificities. These advances may enable the genetically encoded synthesis of non-canonical biopolymers and provide a platform for transforming the discovery and evolution of new materials and therapeutics.

Building genomes to understand biology

Genetic manipulation is one of the central strategies that biologists use to investigate the molecular underpinnings of life and its diversity. Thus, advances in genetic manipulation usually lead to a deeper understanding of biological systems. During the last decade, the construction of chromosomes, known as synthetic genomics, has emerged as a novel approach to genetic manipulation. By facilitating complex modifications to chromosome content and structure, synthetic genomics opens new opportunities for studying biology through genetic manipulation. Here, we discuss different classes of genetic manipulation that are enabled by synthetic genomics, as well as biological problems they each can help solve.

A Lambda Red and FLP/FRT-Mediated Site-Specific Recombination System in Komagataeibacter xylinus and Its Application to Enhance the Productivity of Bacterial Cellulose

Komagataeibacter xylinus has received increasing attention as an important microorganism for the conversion of several carbon sources to bacterial cellulose (BC). However, BC productivity has been impeded by the lack of efficient genetic engineering techniques. In this study, a lambda Red and FLP/FRT-mediated site-specific recombination system was successfully established in Komagataeibacter xylinus. Using this system, the membrane bound gene gcd, a gene that encodes glucose dehydrogenase, was knocked out to reduce the modification of glucose to gluconic acid. The engineered strain could not produce any gluconic acid and presented a decreased bacterial cellulose (BC) production due to its restricted glucose utilization. To address this problem, the gene of glucose facilitator protein (glf; ZMO0366) was introduced into the knockout strain coupled with the overexpression of the endogenous glucokinase gene (glk). The BC yield of the resultant strain increased by 63.63-173.68%, thus reducing the production cost.

λ Recombineering Used to Engineer the Genome of Phage T7

Bacteriophage T7 and T7-like bacteriophages are valuable genetic models for lytic phage biology that have heretofore been intractable with in vivo genetic engineering methods. This manuscript describes that the presence of λ Red recombination proteins makes in vivo recombineering of T7 possible, so that single base changes and whole gene replacements on the T7 genome can be made. Red recombination functions also increase the efficiency of T7 genome DNA transfection of cells by ~100-fold. Likewise, Red function enables two other T7-like bacteriophages that do not normally propagate in E. coli to be recovered following genome transfection. These results constitute major technical advances in the speed and efficiency of bacteriophage T7 engineering and will aid in the rapid development of new phage variants for a variety of applications.

CRISPR-Cas-mediated gene editing in lactic acid bacteria

The high efficiency, convenience and diversity of clustered regular interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) systems are driving a technological revolution in the gene editing of lactic acid bacteria (LAB). Cas-RNA cassettes have been adopted as tools to perform gene deletion, insertion and point mutation in several species of LAB. In this article, we describe the basic mechanisms of the CRISPR-Cas system, and the current gene editing methods available, focusing on the CRISPR-Cas models developed for LAB. We also compare the different types of CRISPR-Cas-based genomic manipulations classified according to the different Cas proteins and the type of recombineering, and discuss the rapidly evolving landscape of CRISPR-Cas application in LAB.

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.

Engineering aminoacyl-tRNA synthetases for use in synthetic biology

Within the broad field of synthetic biology, genetic code expansion (GCE) techniques enable creation of proteins with an expanded set of amino acids. This may be invaluable for applications in therapeutics, bioremediation, and biocatalysis. Central to GCE are aminoacyl-tRNA synthetases (aaRSs) as they link a non-canonical amino acid (ncAA) to their cognate tRNA, allowing ncAA incorporation into proteins on the ribosome. The ncAA-acylating aaRSs and their tRNAs should not cross-react with 20 natural aaRSs and tRNAs in the host, i.e., they need to function as an orthogonal translating system. All current orthogonal aaRS•tRNA pairs have been engineered from naturally occurring molecules to change the aaRS's amino acid specificity or assign the tRNA to a liberated codon of choice. Here we discuss the importance of orthogonality in GCE, laboratory techniques employed to create designer aaRSs and tRNAs, and provide an overview of orthogonal aaRS•tRNA pairs for GCE purposes.

Reporter Phage-Based Detection of Bacterial Pathogens: Design Guidelines and Recent Developments

Fast and reliable detection of bacterial pathogens in clinical samples, contaminated food products, and water supplies can drastically improve clinical outcomes and reduce the socio-economic impact of disease. As natural predators of bacteria, bacteriophages (phages) have evolved to bind their hosts with unparalleled specificity and to rapidly deliver and replicate their viral genome. Not surprisingly, phages and phage-encoded proteins have been used to develop a vast repertoire of diagnostic assays, many of which outperform conventional culture-based and molecular detection methods. While intact phages or phage-encoded affinity proteins can be used to capture bacteria, most phage-inspired detection systems harness viral genome delivery and amplification: to this end, suitable phages are genetically reprogrammed to deliver heterologous reporter genes, whose activity is typically detected through enzymatic substrate conversion to indicate the presence of a viable host cell. Infection with such engineered reporter phages typically leads to a rapid burst of reporter protein production that enables highly sensitive detection. In this review, we highlight recent advances in infection-based detection methods, present guidelines for reporter phage construction, outline technical aspects of reporter phage engineering, and discuss some of the advantages and pitfalls of phage-based pathogen detection. Recent improvements in reporter phage construction and engineering further substantiate the potential of these highly evolved nanomachines as rapid and inexpensive detection systems to replace or complement traditional diagnostic approaches.

Microbial Cell Factory for Efficiently Synthesizing Plant Natural Products via Optimizing the Location and Adaptation of Pathway on Genome Scale

Plant natural products (PNPs) possess important pharmacological activities and are widely used in cosmetics, health care products, and as food additives. Currently, most PNPs are mainly extracted from cultivated plants, and the yield is limited by the long growth cycle, climate change and complex processing steps, which makes the process unsustainable. However, the complex structure of PNPs significantly reduces the efficiency of chemical synthesis. With the development of metabolic engineering and synthetic biology, heterologous biosynthesis of PNPs in microbial cell factories offers an attractive alternative. Based on the in-depth mining and analysis of genome and transcriptome data, the biosynthetic pathways of a number of natural products have been successfully elucidated, which lays the crucial foundation for heterologous production. However, there are several problems in the microbial synthesis of PNPs, including toxicity of intermediates, low enzyme activity, multiple auxotrophic dependence, and uncontrollable metabolic network. Although various metabolic engineering strategies have been developed to solve these problems, optimizing the location and adaptation of pathways on the whole-genome scale is an important strategy in microorganisms. From this perspective, this review introduces the application of CRISPR/Cas9 in editing PNPs biosynthesis pathways in model microorganisms, the influences of pathway location, and the approaches for optimizing the adaptation between metabolic pathways and chassis hosts for facilitating PNPs biosynthesis.

Multiplex gene editing and large DNA fragment deletion by the CRISPR/Cpf1-RecE/T system in Corynebacterium glutamicum

Corynebacterium glutamicum is an essential industrial strain that has been widely harnessed for the production of all kinds of value-added products. Efficient multiplex gene editing and large DNA fragment deletion are essential strategies for industrial biotechnological research. Cpf1 is a robust and simple genome editing tool for simultaneous editing of multiplex genes. However, no studies on effective multiplex gene editing and large DNA fragment deletion by the CRISPR/Cpf1 system in C. glutamicum have been reported. Here, we developed a multiplex gene editing method by optimizing the CRISPR/Cpf1-RecT system and a large chromosomal fragment deletion strategy using the CRISPR/Cpf1-RecET system in C. glutamicum ATCC 14067. The CRISPR/Cpf1-RecT system exhibited a precise editing efficiency of more than 91.6% with the PAM sequences TTTC, TTTG, GTTG or CTTC. The sites that could be edited were limited due to the PAM region and the 1–7 nt at the 5′ end of the protospacer region. Mutations in the PAM region increased the editing efficiency of the − 6 nt region from 0 to 96.7%. Using a crRNA array, two and three genes could be simultaneously edited in one step via the CRISPR/Cpf1-RecT system, and the efficiency of simultaneously editing two genes was 91.6%, but the efficiency of simultaneously editing three genes was below 10%. The editing efficiency for a deletion of 1 kb was 79.6%, and the editing efficiencies for 5- and 20 kb length DNA fragment deletions reached 91.3% and 36.4%, respectively, via the CRISPR/Cpf1-RecET system. This research provides an efficient and simple tool for C. glutamicum genome editing that can further accelerate metabolic engineering efforts and genome evolution.

Mycoplasma pneumoniae Genome Editing Based on Oligo Recombineering and Cas9-Mediated Counterselection

Mycoplasma species share a set of features, such as lack of a cell wall, streamlined genomes, simplified metabolism, and the use of a deviant genetic code, that make them attractive approximations of what a chassis strain should ideally be. Among them, Mycoplasma pneumoniae arises as a candidate for synthetic biology projects, as it is one of the most deeply characterized bacteria. However, the historical paucity of tools for editing Mycoplasma genomes has precluded the establishment of M. pneumoniae as a suitable chassis strain. Here, we developed an oligonucleotide recombineering method for this strain based on GP35, a ssDNA recombinase originally encoded by a Bacillus subtilis-associated phage. GP35-mediated oligo recombineering is able to carry out point mutations in the M. pneumoniae genome with an efficiency as high as 2.7 × 10^–2, outperforming oligo recombineering protocols developed for other bacteria. Gene deletions of different sizes showed a decreasing power trend between efficiency and the scale of the attempted edition. However, the editing rates for all modifications increased when CRISPR/Cas9 was used to counterselect nonedited cells. This allowed edited clones carrying chromosomal deletions of up to 1.8 kb to be recovered with little to no screening of survivor cells. We envision this technology as a major step toward the use of M. pneumoniae, and possibly other Mycoplasmas, as synthetic biology chassis strains.

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.

Non-pathogenic Escherichia coli acquires virulence by mutating a growth-essential LPS transporter

The molecular mechanisms that allow pathogenic bacteria to infect animals have been intensively studied. On the other hand, the molecular mechanisms by which bacteria acquire virulence functions are not fully understood. In the present study, we experimentally evaluated the evolution of a non-pathogenic strain of Escherichia coli in a silkworm infection model and obtained pathogenic mutant strains. As one cause of the high virulence properties of E. coli mutants, we identified amino acid substitutions in LptD (G580S) and LptE (T95I) constituting the lipopolysaccharide (LPS) transporter, which translocates LPS from the inner to the outer membrane and is essential for E. coli growth. The growth of the LptD and LptE mutants obtained in this study was indistinguishable from that of the parent strain. The LptD and LptE mutants exhibited increased secretion of outer membrane vesicles containing LPS and resistance against various antibiotics, antimicrobial peptides, and host complement. In vivo cross-linking studies revealed that the conformation of the LptD-LptE complex was altered in the LptD and LptE mutants. Furthermore, several clinical isolates of E. coli carried amino acid substitutions of LptD and LptE that conferred resistance against antimicrobial substances. This study demonstrated an experimental evolution of bacterial virulence properties in an animal infection model and identified functional alterations of the growth-essential LPS transporter that led to high bacterial virulence by conferring resistance against antimicrobial substances. These findings suggest that non-pathogenic bacteria can gain virulence traits by changing the functions of essential genes, and provide new insight to bacterial evolution in a host environment.

Pathogenic bacteria developed their virulence properties by changing the functions of various genes after the emergence of the host animals on earth. The types of gene function alterations that confer bacterial virulence properties, however, have remained unclear. We utilized a silkworm infection model to perform an experimental evolution of bacterial virulence activity. From a non-pathogenic strain of Escherichia coli, we obtained a mutant strain that exhibited 500-fold higher virulence than the original strain and identified mutations of the lipopolysaccharide (LPS) transporter, which translocates LPS onto the bacterial surface, as one cause of the high virulence. The mutations changed the structure of the LPS transporter, increased the secretion of outer membrane vesicles, and enabled bacterial survival in the presence of host antimicrobial substances. This mechanism to gain high virulence occurs naturally, as several E. coli clinical isolates carried mutations of the LPS transporter that confer resistance against antimicrobial substances. Our study unveiled a novel mechanism by which bacteria increase their virulence through modifying their gene function.

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.

DNA gap repair in Escherichia coli for multiplex site-directed mutagenesis

Site-directed mutagenesis allows the generation of novel DNA sequences that can be used for a variety of important applications such as the functional analysis of genetic variants. To overcome the limitations of existing site-directed mutagenesis approaches, we explored in vivo DNA gap repair. We found that site-specific mutations in plasmid DNA can be generated in Escherichia coli using mutant single-stranded oligonucleotides to target PCR-derived linear double-stranded plasmid DNA. We called this method DeGeRing, and we characterized its advantages, including non-biased multiplex mutagenesis, over existing site-directed mutagenesis methods such as recombineering (recombination-mediated genetic engineering), single DNA break repair (SDBR, introduced by W. Mandecki), and QuikChange (Agilent Technologies, La Jolla, CA). We determined the efficiency of DeGeRing to induce site-directed mutations with and without a phenotype in three K-12 E coli strains using multiple single-stranded oligonucleotides containing homological and heterological parts of various sizes. Virtual lack of background made the isolation of mutants with frequencies up to 10^-6 unnecessary. Our data show that endogenous DNA gap repair in E coli supports efficient multiplex site-directed mutagenesis. DeGeRing might facilitate the generation of mutant DNA sequences for protein engineering and the functional analysis of genetic variants in reverse genetics.

Homing endonuclease I-SceI-mediated Corynebacterium glutamicum ATCC 13032 genome engineering

Corynebacterium glutamicum is widely used to produce amino acids and is a chassis for the production of value-added compounds. Effective genome engineering methods are crucial to metabolic engineering and synthetic biology studies of C. glutamicum. Herein, a homing endonuclease I-SceI-mediated genome engineering strategy was established for the model strain C. glutamicum ATCC 13032. A vegetative R6K replicon-based, suicide plasmid was employed. The plasmid, pLS3661, contains both tightly regulated, IPTG (isopropyl-β-D-1-thiogalactopyranoside)-inducible I-SceI expression elements and two I-SceI recognition sites. Following cloning of the homologous arms into pLS3661 and transfer the recombinant vector into C. glutamicum ATCC 13032, through the homologous recombination between the cloned fragment and its chromosomal allele, a merodiploid was selected under kanamycin selection. Subsequently, a merodiploid was resolved by double-stranded break repair stimulated by IPTG-stimulated I-SceI expression, generating desired mutants. The protocol obviates a pre-generated strain, transfer of a second I-SceI expression plasmid, and there is not any strain, medium, and temperature restrictions. We validated the approach via deletions of five genes (up to ~ 13.0 kb) and knock-in of one DNA fragment. Furthermore, through kanamycin resistance repair, the ssDNA recombineering parameters were optimized. We hope the highly efficient method will be helpful for the studies of C. glutamicum, and potentially, to other bacteria. KEY POINTS: • Counterselection marker I-SceI-mediated C. glutamicum genome engineering • A suicide vector contains I-SceI expression elements and its recognition sites • Gene deletions and knock-in were conducted; efficiency was as high as 90% • Through antibiotic resistance repair, ssDNA recombineering parameters were optimized.

A single digestion, single-stranded oligonucleotide mediated PCR-independent site-directed mutagenesis method

A PCR-independent in vitro site-directed mutagenesis method was established. Cas12a from Francisella novicida (FnCas12a) linearizes the plasmid with single digestion. T5 exonuclease removes the target nucleotide. A short single- or double-stranded mutagenic oligonucleotide introduces the mutation. This rapid and simple mutagenesis method is referred to as FnCas12a and T5 exonuclease mediated site-directed mutagenesis system (CT5-SDM). The platform is also suitable for the mutagenesis of plasmids larger than 10 kb. KEY POINTS: Site-directed mutagenesis mediated by single-stranded DNA. Removing target site with T5 exonuclease. Highly efficient cleavage of target DNA with FnCas12a.

Dynamic Genome Editing Using In Vivo Synthesized Donor ssDNA in Escherichia coli

As a key element of genome editing, donor DNA introduces the desired exogenous sequence while working with other crucial machinery such as CRISPR-Cas or recombinases. However, current methods for the delivery of donor DNA into cells are both inefficient and complicated. Here, we developed a new methodology that utilizes rolling circle replication and Cas9 mediated (RC-Cas-mediated) in vivo single strand DNA (ssDNA) synthesis. A single-gene rolling circle DNA replication system from Gram-negative bacteria was engineered to produce circular ssDNA from a Gram-positive parent plasmid at a designed sequence in Escherichia coli. Furthermore, it was demonstrated that the desired linear ssDNA fragment could be cut out using CRISPR-associated protein 9 (CRISPR-Cas9) nuclease and combined with lambda Red recombinase as donor for precise genome engineering. Various donor ssDNA fragments from hundreds to thousands of nucleotides in length were synthesized in E. coli cells, allowing successive genome editing in growing cells. We hope that this RC-Cas-mediated in vivo ssDNA on-site synthesis system will be widely adopted as a useful new tool for dynamic genome editing.

Multiple-Site Diversification of Regulatory Sequences Enables Interspecies Operability of Genetic Devices

The features of the light-responsive cyanobacterial CcaSR regulatory module that determine interoperability of this optogenetic device between Escherichia coli and Pseudomonas putida have been examined. For this, all structural parts (i.e., ho1 and pcyA genes for synthesis of phycocyanobilin, the ccaS/ccaR system from Synechocystis, and its cognate downstream promoter) were maintained but their expression levels and stoichiometry diversified by (i) reassembling them together in a single broad host range, standardized vector and (ii) subjecting the noncoding regulatory sequences to multiple cycles of directed evolution with random genomic mutations (DIvERGE), a recombineering method that intensifies mutation rates within discrete DNA segments. Once passed to P. putida, various clones displayed a wide dynamic range, insignificant leakiness, and excellent capacity in response to green light. Inspection of the evolutionary intermediates pinpointed translational control as the main bottleneck for interoperability and suggested a general approach for easing the exchange of genetic cargoes between different species, i.e., optimization of relative expression levels and upturning of subcomplex stoichiometry.

Operon Concatenation Is an Ancient Feature That Restricts the Potential to Rearrange Bacterial Chromosomes

The last common ancestor of the Gammaproteobacteria carried an important 40-kb chromosome section encoding 51 proteins of the transcriptional and translational machinery. These genes were organized into eight contiguous operons (rrnB-tufB-secE-rpoBC-str-S10-spc-alpha). Over 2 Gy of evolution, in different lineages, some of the operons became separated by multigene insertions. Surprisingly, in many Enterobacteriaceae, much of the ancient organization is conserved, indicating a strong selective force on the operons to remain colinear. Here, we show for one operon pair, tufB-secE in Salmonella, that an interruption of contiguity significantly reduces growth rate. Our data show that the tufB-secE operons are concatenated by an interoperon terminator–promoter overlap that plays a significant role regulating gene expression. Interrupting operon contiguity interferes with this regulation, reducing cellular fitness. Six operons of the ancestral chromosome section remain contiguous in Salmonella (tufB-secE-rpoBC and S10-spc-alpha) and, strikingly, each of these operon pairs is also connected by an interoperon terminator–promoter overlap. Accordingly, we propose that operon concatenation is an ancient feature that restricts the potential to rearrange bacterial chromosomes and can select for the maintenance of a colinear operon organization over billions of years.

An sRNA Screen for Reversal of Quinolone Resistance in Escherichia coli

In light of the rising prevalence of antimicrobial resistance (AMR) and the slow pace of new antimicrobial development, there has been increasing interest in the development of adjuvants that improve or restore the effectiveness of existing drugs. Here, we use a novel small RNA (sRNA) screening approach to identify genes whose knockdown increases ciprofloxacin (CIP) sensitivity in a resistant strain of Escherichia coli 5000 sRNA constructs were initially screened on a gyrA S83L background, ultimately leading to 30 validated genes whose disruption reduces CIP resistance. This set includes genes involved in DNA replication, repair, recombination, efflux, and other regulatory systems. Our findings increase understanding of the functional interactions of DNA Gyrase, and may aid in the development of new therapeutic approaches for combating AMR.

Recent advances in genetic engineering tools based on synthetic biology

Genome-scale engineering is a crucial methodology to rationally regulate microbiological system operations, leading to expected biological behaviors or enhanced bioproduct yields. Over the past decade, innovative genome modification technologies have been developed for effectively regulating and manipulating genes at the genome level. Here, we discuss the current genome-scale engineering technologies used for microbial engineering. Recently developed strategies, such as clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9, multiplex automated genome engineering (MAGE), promoter engineering, CRISPR-based regulations, and synthetic small regulatory RNA (sRNA)-based knockdown, are considered as powerful tools for genome-scale engineering in microbiological systems. MAGE, which modifies specific nucleotides of the genome sequence, is utilized as a genome-editing tool. Contrastingly, synthetic sRNA, CRISPRi, and CRISPRa are mainly used to regulate gene expression without modifying the genome sequence. This review introduces the recent genome-scale editing and regulating technologies and their applications in metabolic engineering.

SCRaMbLE-in: A Fast and Efficient Method to Diversify and Improve the Yields of Heterologous Pathways in Synthetic Yeast

The synthetic chromosome rearrangement and modification by LoxP-mediated evolution (SCRaMbLE) system is a key component of the synthetic yeast genome (Sc2.0) project, an international effort to construct an entire synthetic genome in yeast. SCRaMbLE involves the introduction of thousands of symmetrical LoxP (LoxPsym) recombination sites downstream of every nonessential gene in all 16 chromosomes, enabling numerous genome rearrangements in the form of deletions, inversions, duplications, and translocations by the Cre-LoxPsym recombination system. We highlight a two-step protocol for SCRaMbLE-in (Liu, Nat Commun 9(1):1936, 2018), a recombinase-based combinatorial method to expedite genetic engineering and exogenous pathway optimization, using a synthetic β-carotene pathway as an example. First, an in vitro phase uses a recombinase toolkit to diversify gene expression by integrating various regulatory elements into the target pathway. This combinatorial pathway library can be transformed directly into yeast for traditional screening. Once an optimized pathway which is flanked by LoxPsym sites is identified, it is transformed into Sc2.0 yeast for the in vivo SCRaMbLE phase, where LoxPsym sites in the synthetic yeast genome and Cre recombinase catalyze massive genome rearrangements. We describe all the conditions necessary to perform SCRaMbLE and post-SCRaMbLE experiments including screening, spot test analysis, and PCRTag analysis to elucidate genotype-phenotype relationships.

A Broad Host Range Plasmid-Based Roadmap for ssDNA-Based Recombineering in Gram-Negative Bacteria

Recombineering is the use of phage recombination proteins to improve and facilitate bacterial genome engineering. Depending on the nature of the DNA template, double-stranded or single-stranded, the system needs three proteins (Gam, Exo, and Beta) or just one (Beta) to work properly. The use of this technique has been fundamental not only toward solving fundamental biological questions with reverse genetics but also for the generation of deep-engineered E. coli chassis strains. Unfortunately, the use of ssDNA recombineering is still limited to a narrow number of bacterial species. One of the reasons for that is the lack of proper recombinases to be efficiently used in different microorganisms and the lack of proper genetic tools to deliver and express this activity in a controlled way. Here, we describe a protocol to follow a simple workflow to identify, clone, and quantify the function of the selected recombinases in the organism of choice by cloning and expressing them in standardized broad host range plasmids. As an example of the method, we tested the use of the Ssr recombinase in P. putida EM42 by introducing a complete deletion of the target gene pyrF. The example shows how two parameters of the mutagenic oligo, i.e., length and phosphorothioate protection, affect the final outcome of the procedure.

Genome Editing in Staphylococcus aureus by Conditional Recombineering and CRISPR/Cas9-Mediated Counterselection

Methods for the genetic manipulation of S. aureus have historically proven challenging, which has hindered experimental studies of this organism. We recently developed a system for recombineering and CRISPR/Cas9-mediated counterselection in S. aureus which utilizes commercially synthesized synthetic DNA oligonucleotides as substrates for introducing precise genomic modifications into the organism and for performing lethal counterselection of unedited cells. These techniques make it possible to scalably and inexpensively engineer desired genomic changes into laboratory or clinical S. aureus strains, using electroporation to introduce the effector plasmid vectors and oligonucleotides. Here we describe detailed protocols for performing genome editing of S. aureus in order to produce isogenic strains using this system and detail general principles which are broadly applicable across a range of organisms for which equivalent systems have been established.

Correction of the aprt Gene Using Repair-Polypurine Reverse Hoogsteen Hairpins in Mammalian Cells

In this study, we describe the correction of single-point mutations in mammalian cells by repair-polypurine reverse Hoogsteen hairpins (repair-PPRHs). These molecules consist of (1) a PPRH hairpin core that binds to a polypyrimidine target sequence in the double-stranded DNA (dsDNA), producing a triplex structure, and (2) an extension sequence homologous to the DNA sequence to be repaired but containing the wild-type nucleotide instead of the mutation and acting as a donor DNA to correct the mutation. We repaired different point mutations in the adenosyl phosphoribosyl transferase (aprt) gene contained in different aprt-deficient Chinese hamster ovary (CHO) cell lines. Because we had previously corrected mutations in the dihydrofolate reductase (dhfr) gene, in this study, we demonstrate the generality of action of the repair-PPRHs. Repaired cells were analyzed by DNA sequencing, mRNA expression, and enzymatic activity to confirm the correction of the mutation. Moreover, whole-genome sequencing analyses did not detect any off-target effect in the repaired genome. We also performed gel-shift assays to show the binding of the repair-PPRH to the target sequence and the formation of a displacement-loop (D-loop) structure that can trigger a homologous recombination event. Overall, we demonstrate that repair-PPRHs achieve the permanent correction of point mutations in the dsDNA at the endogenous level in mammalian cells without off-target activity.

Elements in the λ immunity region regulate phage development: beyond the 'Genetic Switch'

Genetic elements in the bacteriophage λ immunity region contribute to stable maintenance and synchronous induction of the integrated Escherichia coli prophage. There is a bistable switch between lysogenic and lytic growth that is orchestrated by the CI and Cro repressors acting on the lytic (PL and PR ) and lysogenic (PRM ) promoters, referred to as the Genetic Switch. Other less well-characterized elements in the phage immunity region include the PLIT promoter and the immunity terminator, TIMM . The PLIT promoter is repressed by the bacterial LexA protein in λ lysogens. LexA repressor, like the λ CI repressor, is inactivated during the SOS response to DNA damage, and this regulation ensures that the PLIT promoter and the lytic PL and PR promoters are synchronously activated. Proper RexA and RexB protein levels are critical for the switch from lysogeny to lytic growth. Mutation of PLIT reduces RexB levels relative to RexA, compromising cellular energetics and causing a 10-fold reduction in lytic phage yield. The RexA and RexB proteins interact with themselves and each other in a bacterial two-hybrid system. We also find that the transcription terminator, TIMM , is a Rho-independent, intrinsic terminator. Inactivation of TIMM has minimal effect on λ lysogenization or prophage induction.

Tuning chromosomal gene expression in Escherichia coli by combining single-stranded oligonucleotides mediated recombination and kil counter selection system

Extensively modulating gene expression to achieve optimal flux is a critical step in metabolic engineering. Gene expression is usually modulated at the transcriptional level by controlling the strength of a promoter. However, this type of modulation is often hampered by its inability to fully sample the complete continuum of transcriptional control. In Escherichia coli, this limitation can be solved by constructing promoters with a wide range of strengths. In this study, a highly efficient method was developed to modulate a particular chromosomal gene of E. coli at a wide range of expression levels. This was achieved by combining highly efficient single-stranded oligonucleotide-mediated recombination and a stringent counter selection system kil. Using this strategy, a chromosomal library, with a range from 0.3% to 388% relative to the wild lac promoter, was easily obtained. The strength of our chromosomal promoter library was approximately 5-60 times wider in range than those of libraries reported before.

CRISPR/Cas9-enhanced ssDNA recombineering for Pseudomonas putida

Implementation of single-stranded DNA (ssDNA) recombineering in Pseudomonas putida has widened the range of genetic manipulations applicable to this biotechnologically relevant bacterium. Yet, the relatively low efficiency of the technology hampers identification of mutated clones lacking conspicuous phenotypes. Fortunately, the use of CRISPR/Cas9 as a device for counterselection of wild-type sequences helps to overcome this limitation. Merging ssDNA recombineering with CRISPR/Cas9 thus enables a suite of genomic edits with a straightforward approach: a CRISPR plasmid provides the spacer DNA sequence that directs the Cas9 nuclease ribonucleoprotein complex to cleave the genome at the wild-type sequences that have not undergone the change entered by the mutagenic ssDNA oligonucleotide(s). This protocol describes a complete workflow of the method optimized for P. putida, although it could in principle be applicable to many other pseudomonads. As an example, we show the deletion of the edd gene that encodes one key enzyme that operates the EDEMP cycle for glucose metabolism in P. putida EM42. By combining two incompatible CRISPR plasmids with different antibiotic selection markers, we show that the procedure can be cycled to implement consecutive deletions in the same strain, e.g. deletion of the pyrF gene following that of the edd mutant. This approach adds to the wealth of genetic technologies available for P. putida and strengthens its status as a chassis of choice for a suite of biotechnological applications.

Efficient and Precise Genome Editing in Shewanella with Recombineering and CRISPR/Cas9-Mediated Counter-Selection

Dissimilatory metal-reducing bacteria, particularly those from the genus Shewanella, are of importance for bioremediation of metal contaminated sites and sustainable energy production. However, studies on this species have suffered from a lack of effective genetic tools for precise and high throughput genome manipulation. Here we report the development of a highly efficient system based on single-stranded DNA oligonucleotide recombineering coupled with CRISPR/Cas9-mediated counter-selection. Our system uses two plasmids: a sgRNA targeting vector and an editing vector, the latter harboring both Cas9 and the phage recombinase W3 Beta. Following the experimental analysis of Cas9 activity, we demonstrate the ability of this system to efficiently and precisely engineer different Shewanella strains with an average efficiency of >90% among total transformed cells, compared to ≃5% by recombineering alone, and regardless of the gene modified. We also show that different genetic changes can be introduced: mismatches, deletions, and small insertions. Surprisingly, we found that use of CRISPR/Cas9 alone allows selection of recombinase-independent S. oneidensis mutations, albeit at lower efficiency and frequency. With synthesized single-stranded DNA as substrates for homologous recombination and Cas9 as a counter-selectable marker, this new system provides a rapid, scalable, versatile, and scarless tool that will accelerate progress in Shewanella genomic engineering.