Characterizing the portability of phage-encoded homologous recombination proteins

Structural Basis for the Interaction of Redβ Single-Strand Annealing Protein with Escherichia coli Single-Stranded DNA-Binding Protein

Redβ is a protein from bacteriophage λ that binds to single-stranded DNA (ssDNA) to promote the annealing of complementary strands. Together with λ-exonuclease (λ-exo), Redβ is part of a two-component DNA recombination system involved in multiple aspects of genome maintenance. The proteins have been exploited in powerful methods for bacterial genome engineering in which Redβ can anneal an electroporated oligonucleotide to a complementary target site at the lagging strand of a replication fork. Successful annealing in vivo requires the interaction of Redβ with E. coli single-stranded DNA-binding protein (SSB), which coats the ssDNA at the lagging strand to coordinate access of numerous replication proteins. Previous mutational analysis revealed that the interaction between Redβ and SSB involves the C-terminal domain (CTD) of Redβ and the C-terminal tail of SSB (SSB-Ct), the site for binding of numerous host proteins. Here, we have determined the x-ray crystal structure of Redβ CTD in complex with a peptide corresponding to the last nine residues of SSB (MDFDDDIPF). Formation of the complex is predominantly mediated by hydrophobic interactions between two phenylalanine side chains of SSB (Phe-171 and Phe-177) and an apolar groove on the CTD, combined with electrostatic interactions between the C-terminal carboxylate of SSB and Lys-214 of the CTD. Mutation of any of these residues to alanine significantly disrupts the interaction of full-length Redβ and SSB proteins. Structural knowledge of this interaction will help to expand the utility of Redβ-mediated recombination to a wider range of bacterial hosts for applications in synthetic biology.

Genome editing and its role in vaccine, diagnosis, and therapeutic advancement

Genome editing involves precise modification of specific nucleotides in the genome using nucleases like CRISPR/Cas, ZFN, or TALEN, leading to increased efficiency of homologous recombination (HR) for gene editing, and it can result in gene disruption events via non-homologous end joining (NHEJ) or homology-driven repair (HDR). Genome editing, particularly CRISPR-Cas9, revolutionizes vaccine development by enabling precise modifications of pathogen genomes, leading to enhanced vaccine efficacy and safety. It allows for tailored antigen optimization, improved vector design, and deeper insights into host genes' impact on vaccine responses, ultimately enhancing vaccine development and manufacturing processes. This review highlights different types of genome editing methods, their associated risks, approaches to overcome the shortcomings, and the diverse roles of genome editing.

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.

Bacterial genome engineering using CRISPR-associated transposases

Clustered regularly interspaced short palindromic repeats (CRISPR)-associated transposases have the potential to transform the technology landscape for kilobase-scale genome engineering, by virtue of their ability to integrate large genetic payloads with high accuracy, easy programmability and no requirement for homologous recombination machinery. These transposons encode efficient, CRISPR RNA-guided transposases that execute genomic insertions in Escherichia coli at efficiencies approaching ~100%. Moreover, they generate multiplexed edits when programmed with multiple guides, and function robustly in diverse Gram-negative bacterial species. Here we present a detailed protocol for engineering bacterial genomes using CRISPR-associated transposase (CAST) systems, including guidelines on the available vectors, customization of guide RNAs and DNA payloads, selection of common delivery methods, and genotypic analysis of integration events. We further describe a computational CRISPR RNA design algorithm to avoid potential off-targets, and a CRISPR array cloning pipeline for performing multiplexed DNA insertions. The method presented here allows the isolation of clonal strains containing a novel genomic integration event of interest within 1-2 weeks using available plasmid constructs and standard molecular biology techniques.

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.  .

Engineered bacterial orthogonal DNA replication system for continuous evolution

Continuous evolution can generate biomolecules for synthetic biology and enable evolutionary investigation. The orthogonal DNA replication system (OrthoRep) in yeast can efficiently mutate long DNA fragments in an easy-to-operate manner. However, such a system is lacking in bacteria. Therefore, we developed a bacterial orthogonal DNA replication system (BacORep) for continuous evolution. We achieved this by harnessing the temperate phage GIL16 DNA replication machinery in Bacillus thuringiensis with an engineered error-prone orthogonal DNA polymerase. BacORep introduces all 12 types of nucleotide substitution in 15-kilobase genes on orthogonally replicating linear plasmids with a 6,700-fold higher mutation rate than that of the host genome, the mutation rate of which is unchanged. Here we demonstrate the utility of BacORep-based continuous evolution by generating strong promoters applicable to model bacteria, Bacillus subtilis and Escherichia coli, and achieving a 7.4-fold methanol assimilation increase in B. thuringiensis. BacORep is a powerful tool for continuous evolution in prokaryotic cells.

Heterologous Single-Strand DNA-Annealing and Binding Protein Enhance CRISPR-Based Genome Editing Efficiency in Komagataella phaffii

The industrial yeast Komagataella phaffii is a highly effective platform for heterologous protein production, owing to its high protein expression and secretion capacity. Heterologous genes and proteins are involved in multiple processes, including transcription, translation, protein folding, modification, transportation, and degradation; however, engineering these proteins and genes is challenging due to inefficient genome editing techniques. We employed Pseudomonas aeruginosa phage single-stranded DNA-annealing protein (SSAP) PapRecT and P. aeruginosa single-stranded DNA-binding protein (SSB) PaSSB to introduce SSAP-SSB-based homology recombination, which facilitated K. phaffii CRISPR-based genome engineering. Specifically, a host-independent method was developed by expressing sgRNA with PapRecT-PaSSB in a single plasmid, with which only a 50 bp short homologous arm (HA) reached a 100% positive rate for CRISPR-based gene insertion, reaching 18 colony-forming units (CFU) per μg of donor DNA. Single deletion using 1000 bp HA attained 100%, reaching 68 CFUs per μg of donor DNA. Using this efficient CRISPR-based genome editing tool, we integrated three genes (INO4, GAL4-like, and PAB1) at three different loci for overexpression to realize the collaborative regulation of human-lactalbumin (α-LA) production. Specifically, we strengthened phospholipid biosynthesis to facilitate endoplasmic reticulum membrane formation and enhanced recombinant protein transcription and translation by overexpressing transcription and translation factors. The final production of α-LA in the 3 L fermentation reached 113.4 mg L-1, two times higher than that of the strain without multiple site gene editing, which is the highest reported titer in K. phaffii. The CRISPR-based genome editing method developed in this study is suitable for the synergistic multiple-site engineering of protein and biochemical biosynthesis pathways to improve the biomanufacturing efficiency.

A High-Throughput Colocalization Pipeline for Quantification of Mitochondrial Targeting across Different Protein Types

Efficient metabolic engineering and the development of mitochondrial therapeutics often rely upon the specific and strong import of foreign proteins into mitochondria. Fusing a protein to a mitochondria-bound signal peptide is a common method to localize proteins to mitochondria, but this strategy is not universally effective, with particular proteins empirically failing to localize. To help overcome this barrier, this work develops a generalizable and open-source framework to design proteins for mitochondrial import and quantify their specific localization. This Python-based pipeline quantitatively assesses the colocalization of different proteins previously used for precise genome editing in a high-throughput manner to reveal signal peptide-protein combinations that localize well in mitochondria.

Genetic engineering of bacteriophages: Key concepts, strategies, and applications

Bacteriophages are the most abundant biological entity in the world and hold a tremendous amount of unexplored genetic information. Since their discovery, phages have drawn a great deal of attention from researchers despite their small size. The development of advanced strategies to modify their genomes and produce engineered phages with desired traits has opened new avenues for their applications. This review presents advanced strategies for developing engineered phages and their potential antibacterial applications in phage therapy, disruption of biofilm, delivery of antimicrobials, use of endolysin as an antibacterial agent, and altering the phage host range. Similarly, engineered phages find applications in eukaryotes as a shuttle for delivering genes and drugs to the targeted cells, and are used in the development of vaccines and facilitating tissue engineering. The use of phage display-based specific peptides for vaccine development, diagnostic tools, and targeted drug delivery is also discussed in this review. The engineered phage-mediated industrial food processing and biocontrol, advanced wastewater treatment, phage-based nano-medicines, and their use as a bio-recognition element for the detection of bacterial pathogens are also part of this review. The genetic engineering approaches hold great potential to accelerate translational phages and research. Overall, this review provides a deep understanding of the ingenious knowledge of phage engineering to move them beyond their innate ability for potential applications.

High-throughput colocalization pipeline quantifies efficacy of mitochondrial targeting signals across different protein types

Efficient metabolic engineering and the development of mitochondrial therapeutics often rely upon the specific and strong import of foreign proteins into mitochondria. Fusing a protein to a mitochondria-bound signal peptide is a common method to localize proteins to mitochondria, but this strategy is not universally effective with particular proteins empirically failing to localize. To help overcome this barrier, this work develops a generalizable and open-source framework to design proteins for mitochondrial import and quantify their specific localization. By using a Python-based pipeline to quantitatively assess the colocalization of different proteins previously used for precise genome editing in a high-throughput manner, we reveal signal peptide-protein combinations that localize well in mitochondria and, more broadly, general trends about the overall reliability of commonly used mitochondrial targeting signals.

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.

Bacterial genome engineering using CRISPR RNA-guided transposases

CRISPR-associated transposons (CASTs) have the potential to transform the technology landscape for kilobase-scale genome engineering, by virtue of their ability to integrate large genetic payloads with high accuracy, easy programmability, and no requirement for homologous recombination machinery. These transposons encode efficient, CRISPR RNA-guided transposases that execute genomic insertions in E. coli at efficiencies approaching ~100%, generate multiplexed edits when programmed with multiple guides, and function robustly in diverse Gram-negative bacterial species. Here we present a detailed protocol for engineering bacterial genomes using CAST systems, including guidelines on the available homologs and vectors, customization of guide RNAs and DNA payloads, selection of common delivery methods, and genotypic analysis of integration events. We further describe a computational crRNA design algorithm to avoid potential off-targets and CRISPR array cloning pipeline for DNA insertion multiplexing. Starting from available plasmid constructs, the isolation of clonal strains containing a novel genomic integration event-of-interest can be achieved in 1 week using standard molecular biology techniques.

Recombineering-Mediated Sinorhizobium meliloti Rm1021 Gene Deletion

Sinorhizobium meliloti Rm1021 (S. meliloti Rm1021) is a Gram-negative, soil-dwelling α-proteobacterium which serves as a model microorganism for the studies of symbiotic nitrogen fixation. The S. meliloti Rm1021 genome consists of one chromosome and two megaplasmids, pSymA and pSymB. Gene deletion is an essential tool for the elucidation of gene function and generation of mutants with improved properties. However, only two gene deletion methods, counterselectable marker sacB-based and FLP/FRT, Cre/loxP site-specific recombination, have been reported for S. meliloti Rm1021 gene deletion. Both methods require time-consuming and tedious gene cloning and conjugation steps. Herein, a λ Red recombineering-mediated gene deletion strategy is reported. The mutant was obtained via electroporating overlap-extension PCR-generated linear targeting DNA into Red-proficient cells. One gene each from the S. meliloti Rm1021 chromosome, megaplasmid SymA and pSymB was deleted, with deletion efficiency up to 100%. The straightforward and highly efficient recombineering procedure holds the promise to be a general gene manipulation method for S. meliloti Rm1021.

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.

Editing of Phage Genomes—Recombineering-assisted SpCas9 Modification of Model Coliphages T7, T5, and T3

Abstract

Bacteriophages—viruses that infect bacterial cells—are the most abundant biological entities on Earth. The use of phages in fundamental research and industry requires tools for precise manipulation of their genomes. Yet, compared to bacterial genome engineering, modification of phage genomes is challenging because of the lack of selective markers and thus requires laborious screenings of recombinant/mutated phage variants. The development of the CRISPR-Cas technologies allowed to solve this issue by the implementation of negative selection that eliminates the parental phage genomes. In this manuscript, we summarize current methods of phage genome engineering and their coupling with CRISPR-Cas technologies. We also provide examples of our successful application of these methods for introduction of specific insertions, deletions, and point mutations in the genomes of model Escherichia coli lytic phages T7, T5, and T3.

A multifunctional system for genome editing and large-scale interspecies gene transfer

CRISPR SWAPnDROP extends the limits of genome editing to large-scale in-vivo DNA transfer between bacterial species. Its modular platform approach facilitates species specific adaptation to confer genome editing in various species. In this study, we show the implementation of the CRISPR SWAPnDROP concept for the model organism Escherichia coli, the fast growing Vibrio natriegens and the plant pathogen Dickeya dadantii. We demonstrate the excision, transfer and integration of large chromosomal regions between E. coli, V. natriegens and D. dadantii without size-limiting intermediate DNA extraction. CRISPR SWAPnDROP also provides common genome editing approaches comprising scarless, marker-free, iterative and parallel insertions and deletions. The modular character facilitates DNA library applications, and recycling of standardized parts. Its multi-color scarless co-selection system significantly improves editing efficiency and provides visual quality controls throughout the assembly and editing process.

Genome Engineering in Mycoplasma gallisepticum Using Exogenous Recombination Systems

Mycoplasma gallisepticum (Mgal) is a common pathogen of poultry worldwide that has recently spread to North American house finches after a single host shift in 1994. The molecular determinants of Mgal virulence and host specificity are still largely unknown, mostly due to the absence of efficient methods for functional genomics. After evaluating two exogenous recombination systems derived from phages found in the phylogenetically related Spiroplasma phoeniceum and the more distant Bacillus subtilis, the RecET-like system from B. subtilis was successfully used for gene inactivation and targeted replacement in Mgal. In a second step, the Cre-lox recombination system was used for the removal of the antibiotic resistance marker in recombinant mutants. This study therefore describes the first genetic tool for targeted genome engineering of Mgal and demonstrates the efficiency of heterologous recombination systems in minimal bacteria.

A Mini-Review of Flavone Isomers Apigenin and Genistein in Prostate Cancer Treatment

The initial responses to standard chemotherapies among prostate cancer (PCa) patients are usually significant, while most of them will finally develop drug resistance, rendering them with limited therapies. To discover new regimens for the treatment of PCa including resistant PCa, natural products, the richest source of bioactive compounds, can serve as a library for screening and identifying promising candidates, and flavones such as apigenin and genistein have been used in lab and clinical trials for treating PCa over decades. In this mini-review, we take a look into the progress of apigenin and genistein, which are isomers, in treating PCa in the past decade. While possessing very similar structure, these two isomers can both target the same signaling pathways; they also are found to work differently in PCa cells. Given that more combinations are being developed and tested, genistein appears to be the more promising option to be approved. The anticancer efficacies of these two flavones can be confirmed by in-vitro and in-vivo studies, and their applications remain to be validated in clinical trials. Information gained in this work may provide important information for new drug development and the potential application of apigenin and genistein in treating PCa.

CRISPR-based genome editing through the lens of DNA repair

Genome editing technologies operate by inducing site-specific DNA perturbations that are resolved by cellular DNA repair pathways. Products of genome editors include DNA breaks generated by CRISPR-associated nucleases, base modifications induced by base editors, DNA flaps created by prime editors, and integration intermediates formed by site-specific recombinases and transposases associated with CRISPR systems. Here, we discuss the cellular processes that repair CRISPR-generated DNA lesions and describe strategies to obtain desirable genomic changes through modulation of DNA repair pathways. Advances in our understanding of the DNA repair circuitry, in conjunction with the rapid development of innovative genome editing technologies, promise to greatly enhance our ability to improve food production, combat environmental pollution, develop cell-based therapies, and cure genetic and infectious diseases.

Resolving the structure of phage-bacteria interactions in the context of natural diversity

Microbial communities are shaped by viral predators. Yet, resolving which viruses (phages) and bacteria are interacting is a major challenge in the context of natural levels of microbial diversity. Thus, fundamental features of how phage-bacteria interactions are structured and evolve in the wild remain poorly resolved. Here we use large-scale isolation of environmental marine Vibrio bacteria and their phages to obtain estimates of strain-level phage predator loads, and use all-by-all host range assays to discover how phage and host genomic diversity shape interactions. We show that lytic interactions in environmental interaction networks (as observed in agar overlay) are sparse-with phage predator loads being low for most bacterial strains, and phages being host-strain-specific. Paradoxically, we also find that although overlap in killing is generally rare between tailed phages, recombination is common. Together, these results suggest that recombination during cryptic co-infections is an important mode of phage evolution in microbial communities. In the development of phages for bioengineering and therapeutics it is important to consider that nucleic acids of introduced phages may spread into local phage populations through recombination, and that the likelihood of transfer is not predictable based on lytic host range.

DNA Damage Protection for Enhanced Bacterial Survival Under Simulated Low Earth Orbit Environmental Conditions in Escherichia coli

Some organisms have shown the ability to naturally survive in extreme environments, even outer space. Some of these have natural mechanisms to resist severe DNA damage from conditions such as ionizing and non-ionizing radiation, extreme temperatures, and low pressures or vacuum. A good example can be found in Deinococcus radiodurans, which was exposed to severe conditions such as those listed in the Exposure Facility of the International Space Station (ISS) for up to three years. Another example are tardigrades (Ramazzottius varieornatus) which are some of the most resilient animals known. In this study, the survival under simulated Low earth Orbit (LEO) environmental conditions was tested in Escherichia coli. The radiation resistance of this bacteria was enhanced using the Dsup gene from R. varieornatus, and two more genes from D. radiodurans involved in DNA damage repair, RecA and uvrD. The enhanced survival to wide ranges of temperatures and low pressures was then tested in the new strains. This research constitutes a first step in the creation of new bacterial strains engineered to survive severe conditions and adapting existing species for their survival in remote environments, including extra-terrestrial habitats. These strains could be key for the development of environments hospitable to life and could be of use for ecological restoration and space exploration. In addition, studying the efficacy and the functioning of the DNA repair mechanisms used in this study could be beneficial for medical and life sciences engineering.

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

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

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

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

A CRISPR/Cas9-based single-stranded DNA recombineering system for genome editing of Rhodococcus opacus PD630

Genome engineering of Rhodococcus opacus PD630, an important microorganism used for the bioconversion of lignin, is currently dependent on inefficient homologous recombination. Although a CRISPR interference procedure for gene repression has previously been developed for R. opacus PD630, a CRISPR/Cas9 system for gene knockout has yet to be reported for the strain. In this study, we found that the cytotoxicity of Cas9 and the deficiency in pathways for repairing DNA double-strand breaks (DSBs) were the major causes of the failure of conventional CRISPR/Cas9 technologies in R. opacus, even when augmented with the recombinases Che9c60 and Che9c61. We successfully developed an efficient single-stranded DNA (ssDNA) recombineering system coupled with CRISPR/Cas9 counter-selection, which facilitated rapid and scarless editing of the R. opacus genome. A two-plasmid system, comprising Cas9 driven by a weak Rhodococcus promoter Pniami, designed to prevent cytotoxicity, and a single-guide RNA (sgRNA) under the control of a strong constitutive promoter, was proven to be appropriate with respect to cleavage function. A novel recombinase, RrRecT derived from a Rhodococcus ruber prophage, was identified for the first time, which facilitated recombination of short ssDNA donors (40–80 nt) targeted to the lagging strand and enabled us to obtain a recombination efficiency up to 10³-fold higher than that of endogenous pathways. Finally, by incorporating RrRecT and Cas9 into a single plasmid and then co-transforming cells with sgRNA plasmids and short ssDNA donors, we efficiently achieved gene disruption and base mutation in R. opacus, with editing efficiencies ranging from 22 % to 100 %. Simultaneous disruption of double genes was also confirmed, although at a lower efficiency. This effective genome editing tool will accelerate the engineering of R. opacus metabolism.

RecT Recombinase Expression Enables Efficient Gene Editing in Enterococcus spp

Enterococcus faecium is a ubiquitous Gram-positive bacterium that has been recovered from the environment, food, and microbiota of mammals. Commensal strains of E. faecium can confer beneficial effects on host physiology and immunity, but antibiotic usage has afforded antibiotic-resistant and pathogenic isolates from livestock and humans. However, the dissection of E. faecium functions and mechanisms has been restricted by inefficient gene-editing methods. To address these limitations, here, we report that the expression of E. faecium RecT recombinase significantly improves the efficiency of recombineering technologies in both commensal and antibiotic-resistant strains of E. faecium and other Enterococcus species such as E. durans and E. hirae. Notably, the expression of RecT in combination with clustered regularly interspaced short palindromic repeat (CRISPR)-Cas9 and guide RNAs (gRNAs) enabled highly efficient scarless single-stranded DNA recombineering to generate specific gene-editing mutants in E. faecium. Moreover, we demonstrate that E. faecium RecT expression facilitated chromosomal insertions of double-stranded DNA templates encoding antibiotic-selectable markers to generate gene deletion mutants. As a further proof of principle, we use CRISPR-Cas9-mediated recombineering to knock out both sortase A genes in E. faecium for downstream functional characterization. The general RecT-mediated recombineering methods described here should significantly enhance genetic studies of E. faecium and other closely related species for functional and mechanistic studies. IMPORTANCE Enterococcus faecium is widely recognized as an emerging public health threat with the rise of drug resistance and nosocomial infections. Nevertheless, commensal Enterococcus strains possess beneficial health functions in mammals to upregulate host immunity and prevent microbial infections. This functional dichotomy of Enterococcus species and strains highlights the need for in-depth studies to discover and characterize the genetic components underlying its diverse activities. However, current genetic engineering methods in E. faecium still require passive homologous recombination from plasmid DNA. This involves the successful cloning of multiple homologous fragments into a plasmid, introducing the plasmid into E. faecium, and screening for double-crossover events that can collectively take up to multiple weeks to perform. To alleviate these challenges, we show that RecT recombinase enables the rapid and efficient integration of mutagenic DNA templates to generate substitutions, deletions, and insertions in the genomic DNA of E. faecium. These improved recombineering methods should facilitate functional and mechanistic studies of Enterococcus.

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

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

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.