Single-stranded heteroduplex intermediates in lambda Red homologous recombination

ReaL-MGE is a tool for enhanced multiplex genome engineering and application to malonyl-CoA anabolism

The complexities encountered in microbial metabolic engineering continue to elude prediction and design. Unravelling these complexities requires strategies that go beyond conventional genetics. Using multiplex mutagenesis with double stranded (ds) DNA, we extend the multiplex repertoire previously pioneered using single strand (ss) oligonucleotides. We present ReaL-MGE (Recombineering and Linear CRISPR/Cas9 assisted Multiplex Genome Engineering). ReaL-MGE enables precise manipulation of numerous large DNA sequences as demonstrated by the simultaneous insertion of multiple kilobase-scale sequences into E. coli, Schlegelella brevitalea and Pseudomonas putida genomes without any off-target errors. ReaL-MGE applications to enhance intracellular malonyl-CoA levels in these three genomes achieved 26-, 20-, and 13.5-fold elevations respectively, thereby promoting target polyketide yields by more than an order of magnitude. In a further round of ReaL-MGE, we adapt S. brevitalea to malonyl-CoA elevation utilizing a restricted carbon source (lignocellulose from straw) to realize production of the anti-cancer secondary metabolite, epothilone from lignocellulose. Multiplex mutagenesis with dsDNA enables the incorporation of lengthy segments that can fully encode additional functions. Additionally, the utilization of PCR to generate the dsDNAs brings flexible design advantages. ReaL-MGE presents strategic options in microbial metabolic engineering.

Proof of ssDNA degraded from dsDNA for ET recombination

The widely used ET recombination requires an ssDNA product degraded by Rac phage protein E588 from dsDNA for strand invasion. However, proof of the ssDNA product is still elusive. The study provided three levels of proof sequentially. The probable ssDNAs degraded by E588 from the fluorescent plus-, minus-, or double-stranded dsDNA pET28a-xylanase exhibited a half fluorescence intensity of the corresponding dsDNAs, equivalent to the E588 degradation nucleotides half that of the total nucleotides degraded from the corresponding dsDNA. The ssDNA product degraded by E588 from the fluorescent minus-stranded dsDNA was confirmed by gradient gel-electrophoresis and two nuclease degradation reactions. Degraded by E588 from the dsDNA pET28a-xylanase that had a phosphorothioated plus-stranded 5'-terminus, the plus-stranded ssDNA product was separated via gel electrophoresis and recovered via a DNAclean kit. The recovered ssDNA product was proven to have intact 5'- and 3'-ends by DNA sequencing analysis. This study provides a solid foundation for the mechanism of ssDNA invasion.

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.

DNA Recombineering Applications

The ability to manipulate the bacterial genome is an obligatory premise for the study of gene function and regulation in bacterial cells. The λ red recombineering technique allows modification of chromosomal sequences with base-pair precision without the need of intermediate molecular cloning steps. Initially conceived to construct insertion mutants, the technique lends itself to a wide variety of applications including the creation of point mutants, seamless deletions, reporter, and epitope tag fusions and chromosomal rearrangements. Here, we introduce some of the most common implementations of the method.

High production of poly(3-hydroxybutyrate) in Escherichia coli using crude glycerol

Poly(3-hydroxybutyrate) (PHB) is a prominent bio-plastic and recognized as the potential replacement of petroleum-derived plastics. To make PHB cost-effective, the production scheme based on crude glycerol was developed using Escherichia coli. The heterogeneous synthesis pathway of PHB was introduced into the E. coli strain capable of efficiently utilizing glycerol. The central metabolism that links to the synthesis of acetyl-CoA and NADPH was further reprogrammed to improve the PHB production. Key genes were targeted for manipulation, involving those in glycolysis, the pentose phosphate pathway, and the tricarboxylic cycle. As a result, the engineered strain gained a 22-fold increase in the PHB titer. Finally, the fed-batch fermentation was conducted with the producer strain to give the PHB titer, content, and productivity reaching 36.3±3.0 g/L, 66.5±2.8%, and 1.2±0.1 g/L/h, respectively. The PHB yield on crude glycerol accounts for 0.3 g/g. The result indicates that the technology platform as developed is promising for the production of bio-plastics.

Single-Stranded DNA-Binding Proteins Mediate DSB Repair and Effectively Improve CRISPR/Cas9 Genome Editing in Escherichia coli and Pseudomonas

Single-stranded DNA-binding proteins (SSBs) are essential for all living organisms. Whether SSBs can repair DNA double-strand breaks (DSBs) and improve the efficiency of CRISPR/Cas9-mediated genome editing has not been determined. Here, based on a pCas/pTargetF system, we constructed pCas-SSB and pCas-T4L by replacing the λ-Red recombinases with Escherichia coli SSB and phage T4 DNA ligase in pCas, respectively. Inactivation of the E. coli lacZ gene with homologous donor dsDNA increased the gene editing efficiency of pCas-SSB/pTargetF by 21.4% compared to pCas/pTargetF. Inactivation of the E. coli lacZ gene via NHEJ increased the gene editing efficiency of pCas-SSB/pTargetF by 33.2% compared to pCas-T4L/pTargetF. Furthermore, the gene-editing efficiency of pCas-SSB/pTargetF in E. coli (ΔrecA, ΔrecBCD, ΔSSB) with or without donor dsDNA did not differ. Additionally, pCas-SSB/pTargetF with donor dsDNA successfully deleted the wp116 gene in Pseudomonas sp. UW4. These results demonstrate that E. coli SSB repairs DSBs caused by CRISPR/Cas9 and effectively improves CRISPR/Cas9 genome editing in E. coli and Pseudomonas.

A Toolkit for Effective and Successive Genome Engineering of Escherichia coli

The bacterium Escherichia coli has been well-justified as an effective workhorse for industrial applications. In this study, we developed a toolkit for flexible genome engineering of this microorganism, including site-specific insertion of heterologous genes and inactivation of endogenous genes, such that bacterial hosts can be effectively engineered for biomanufacturing. We first constructed a base strain by genomic implementation of the cas9 and λRed recombineering genes. Then, we constructed plasmids for expressing gRNA, DNA cargo, and the Vibrio cholerae Tn6677 transposon and type I-F CRISPR-Cas machinery. Genomic insertion of a DNA cargo up to 5.5 kb was conducted using a transposon-associated CRISPR-Cas system, whereas gene inactivation was mediated by a classic CRISPR-Cas9 system coupled with λRed recombineering. With this toolkit, we can exploit the synergistic functions of CRISPR-Cas, λRed recombineering, and Tn6677 transposon for successive genomic manipulations. As a demonstration, we used the developed toolkit to derive a plasmid-free strain for heterologous production of poly(3-hydroxybutyrate-co-3-hydroxyvalerate) (PHBV) by genomic knock-in and knockout of several key genes with high editing efficiencies.

Redβ177 annealase structure reveals details of oligomerization and λ Red-mediated homologous DNA recombination

The Redβ protein of the bacteriophage λ red recombination system is a model annealase which catalyzes single-strand annealing homologous DNA recombination. Here we present the structure of a helical oligomeric annealing intermediate of Redβ, consisting of N-terminal residues 1-177 bound to two complementary 27mer oligonucleotides, determined via cryogenic electron microscopy (cryo-EM) to a final resolution of 3.3 Å. The structure reveals a continuous binding groove which positions and stabilizes complementary DNA strands in a planar orientation to facilitate base pairing via a network of hydrogen bonding. Definition of the inter-subunit interface provides a structural basis for the propensity of Redβ to oligomerize into functionally significant long helical filaments, a trait shared by most annealases. Our cryo-EM structure and molecular dynamics simulations suggest that residues 133-138 form a flexible loop which modulates access to the binding groove. More than half a century after its discovery, this combination of structural and computational observations has allowed us to propose molecular mechanisms for the actions of the model annealase Redβ, a defining member of the Redβ/RecT protein family.

Redβ annealase catalyses single-strand annealing homologous DNA recombination. Here, the authors present a cryo-EM structure of a Redβ annealing intermediate bound to two complementary 27mer oligonucleotides.

Robust counterselection and advanced λRed recombineering enable markerless chromosomal integration of large heterologous constructs

Despite advances in bacterial genome engineering, delivery of large synthetic constructs remains challenging in practice. In this study, we propose a straightforward and robust approach for the markerless integration of DNA fragments encoding whole metabolic pathways into the genome. This approach relies on the replacement of a counterselection marker with cargo DNA cassettes via λRed recombineering. We employed a counterselection strategy involving a genetic circuit based on the CI repressor of λ phage. Our design ensures elimination of most spontaneous mutants, and thus provides a counterselection stringency close to the maximum possible. We improved the efficiency of integrating long PCR-generated cassettes by exploiting the Ocr antirestriction function of T7 phage, which completely prevents degradation of unmethylated DNA by restriction endonucleases in wild-type bacteria. The employment of highly restrictive counterselection and ocr-assisted λRed recombineering allowed markerless integration of operon-sized cassettes into arbitrary genomic loci of four enterobacterial species with an efficiency of 50–100%. In the case of Escherichia coli, our strategy ensures simple combination of markerless mutations in a single strain via P1 transduction. Overall, the proposed approach can serve as a general tool for synthetic biology and metabolic engineering in a range of bacterial hosts.

Cytolysin A-mediated protein exportation efficiency and its role in enhancing the fitness of live recombinant Salmonella Typhi vaccine strain

The genetic fusion of cytolysin A (ClyA) to heterologous antigen expressed in live Salmonella vector demonstrated efficient translocation into periplasmic space and extracellular medium. Accumulating evidence has shown that clyA-mediated antigen delivery improved growth fitness and enhanced immunogenicity of live vector vaccine, but the factors influencing this protein exportation has not been investigated. In this study, Toxoplasma gondii antigen fused at C-terminal of clyA protein was expressed in live S. Typhi vector via both plasmid and chromosomal-based expressions. The bivalent strains showed comparable growth rates as monovalent strains, but in varies antigen exportation efficiency. ClyA-fusion antigen with positive charges were translocated to the extracellular spaces, whereas those with negative charges were retained in the cytoplasm. Furthermore, excessive cellular resources expenditure on antigen expression, especially antigen with larger size, could limit the clyA-fusion antigen exportation, resulting in undesirable metabolic burden that eventually affects the growth fitness. Altogether, the present work indicates potential linkage of factors mainly on antigen properties and expression platforms that may affect clyA-mediated antigen delivery to enhance the growth fitness of live vector strain.

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

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

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.

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.

Half a century of bacteriophage lambda recombinase: In vitro studies of lambda exonuclease and Red-beta annealase

DNA recombination, replication, and repair are intrinsically interconnected processes. From viruses to humans, they are ubiquitous and essential to all life on Earth. Single‐strand annealing homologous DNA recombination is a major mechanism for the repair of double‐stranded DNA breaks. An exonuclease and an annealase work in tandem, forming a complex known as a two‐component recombinase. Redβ annealase and λ‐exonuclease from phage lambda form the archetypal two‐component recombinase complex. In this short review article, we highlight some of the in vitro studies that have led to our current understanding of the lambda recombinase system. We synthesize insights from more than half a century of research, summarizing the state of our current understanding. From this foundation, we identify the gaps in our knowledge and cast an eye forward to consider what the next 50 years of research may uncover.

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.

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

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

Recombineering for Genetic Engineering of Natural Product Biosynthetic Pathways

Microbial genomes encode many cryptic and uncharacterized biosynthetic gene clusters (BGCs). Exploiting this unexplored genetic wealth to discover microbial novel natural products (NPs) remains a challenging issue. We review homologous recombination (HR)-based recombineering, mediated by the recombinases RecE/RecT from Rac prophage and Redα/Redβ from lambda phage, which has developed into a highly inclusive tool for direct cloning of large DNA up to 100 kb, seamless mutation, multifragment assembly, and heterologous expression of microbial NP BGCs. Its utilization in the refactoring, engineering, and functional expression of long BGCs for NP biosynthesis makes it easy to elucidate NP-producing potential in microbes. This review also highlights various applications of recombineering in NP-derived drug discovery.

Nucleocapsid Assembly of Baculoviruses

The baculovirus nucleocapsid is formed through a rod-like capsid encapsulating a genomic DNA molecule of 80~180 kbp. The viral capsid is a large oligomer composed of many copies of various protein subunits. The assembly of viral capsids is a complex oligomerization process. The timing of expression of nucleocapsid-related proteins, transport pathways, and their interactions can affect the assembly process of preformed capsids. In addition, the selection of viral DNA and the injection of the viral genome into empty capsids are the critical steps in nucleocapsid assembly. This paper reviews the replication and recombination of baculovirus DNA, expression and transport of capsid proteins, formation of preformed capsids, DNA encapsulation, and nucleocapsid formation. This review will provide a basis for further study of the nucleocapsid assembly mechanism of baculovirus.

Structure and mechanism of the Red recombination system of bacteriophage λ

While much of this volume focuses on mammalian DNA repair systems that are directly involved in genome stability and cancer, it is important to still be mindful of model systems from prokaryotes. Herein we review the Red recombination system of bacteriophage λ, which consists of an exonuclease for resecting dsDNA ends, and a single-strand annealing protein (SSAP) for binding the resulting 3'-overhang and annealing it to a complementary strand. The genetics and biochemistry of Red have been studied for over 50 years, in work that has laid much of the foundation for understanding DNA recombination in higher eukaryotes. In fact, the Red exonuclease (λ exo) is homologous to Dna2, a nuclease involved in DNA end-resection in eukaryotes, and the Red annealing protein (Redβ) is homologous to Rad52, the primary SSAP in eukaryotes. While eukaryotic recombination involves an elaborate network of proteins that is still being unraveled, the phage systems are comparatively simple and streamlined, yet still encompass the fundamental features of recombination, namely DNA end-resection, homologous pairing (annealing), and a coupling between them. Moreover, the Red system has been exploited in powerful methods for bacterial genome engineering that are important for functional genomics and systems biology. However, several mechanistic aspects of Red, particularly the action of the annealing protein, remain poorly understood. This review will focus on the proteins of the Red recombination system, with particular attention to structural and mechanistic aspects, and how the lessons learned can be applied to eukaryotic systems.

Single-Stranded DNA-Binding Protein and Exogenous RecBCD Inhibitors Enhance Phage-Derived Homologous Recombination in Pseudomonas

The limited efficiency of the available tools for genetic manipulation of Pseudomonas limits fundamental research and utilization of this genus. We explored the properties of a lambda Red-like operon (BAS) from Pseudomonas aeruginosa phage Ab31 and a Rac bacteriophage RecET-like operon (RecTE_Psy) from Pseudomonas syringae pv. syringae B728a. Compared with RecTE_Psy, the BAS operon was functional at a higher temperature indicating potential to be a generic system for Pseudomonas. Owing to the lack of RecBCD inhibitor in the BAS operon, we added Redγ or Pluγ and found increased recombineering efficiencies in P. aeruginosa and Pseudomonas fluorescens but not in Pseudomonas putida and P. syringae. Overexpression of single-stranded DNA-binding protein enhanced recombineering in several contexts including RecET recombination in E. coli. The utility of these systems was demonstrated by engineering P. aeruginosa genomes to create an attenuated rhamnolipid producer. Our work enhances the potential for functional genomics in Pseudomonas.

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The BAS operon is a generic recombineering system for Pseudomonas species

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Single-stranded DNA-binding proteins (SSBs) can stimulate homologous recombination

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The heterologous gam genes can inhibit RecBCD function in Pseudomonas

Excision of selectable markers from the Escherichia coli genome without counterselection using an optimized λRed recombineering procedure

The introduction of chromosomal mutations into the E. coli genome using λRed-mediated recombineering includes two consecutive steps-the insertion of an antibiotic resistance gene and the subsequent excision of the marker. The second step usually requires a counterselection method, because the efficiency of recombination is not high enough to find recombinants among non-recombinant cells. Most counterselection methods require the introduction of additional mutations into the genome or the use of expensive chemicals. In this paper, we describe the development of a reliable procedure for the removal of an antibiotic resistance marker from the E. coli genome without the need for counterselection. For this purpose, we used dsDNA cassettes consisting of two regions homologous to the sequences that flank the marker on the chromosome. We optimized the length of the homologous regions, the electroporation conditions, and the duration of recovery for the electroporated cells in order to maximize the recombination efficiency. Using the optimal parameters identified, we obtained a rate of 4-6% recombinants among the transformed cells. This high efficiency allowed us to find marker-less, antibiotic-sensitive recombinants by replica plating without the need for selection.

Herpes simplex virus 1 ICP8 mutant lacking annealing activity is deficient for viral DNA replication

Most DNA viruses that use recombination-dependent mechanisms to replicate their DNA encode a single-strand annealing protein (SSAP). The herpes simplex virus (HSV) single-strand DNA binding protein (SSB), ICP8, is the central player in all stages of DNA replication. ICP8 is a classical replicative SSB and interacts physically and/or functionally with the other viral replication proteins. Additionally, ICP8 can promote efficient annealing of complementary ssDNA and is thus considered to be a member of the SSAP family. The role of annealing during HSV infection has been difficult to assess in part, because it has not been possible to distinguish between the role of ICP8 as an SSAP from its role as a replicative SSB during viral replication. In this paper, we have characterized an ICP8 mutant, Q706A/F707A (QF), that lacks annealing activity but retains many other functions characteristic of replicative SSBs. Like WT ICP8, the QF mutant protein forms filaments in vitro, binds ssDNA cooperatively, and stimulates the activities of other replication proteins including the viral polymerase, helicase-primase complex, and the origin binding protein. Interestingly, the QF mutant does not complement an ICP8-null virus for viral growth, replication compartment formation, or DNA replication. Thus, we have been able to separate the activities of ICP8 as a replicative SSB from its annealing activity. Taken together, our data indicate that the annealing activity of ICP8 is essential for viral DNA replication in the context of infection and support the notion that HSV-1 uses recombination-dependent mechanisms during DNA replication.

Development of new versatile plasmid-based systems for λRed-mediated Escherichia coli genome engineering

Plasmid-based systems are the most appropriate for multistep lambda Red (λRed)-mediated recombineering, such as the assembly of strains for biotechnological applications. Currently, the widely used λRed-expressing plasmids use a temperature-sensitive origin of replication or temperature shift control of λRed expression. In this work, we have constructed a new, conditionally replicating vector that can be efficiently eliminated from the host strain through passaging in medium containing isopropyl-β-d-thiogalactopyranoside. Using the new vector, we have developed two improved helper plasmids (viz., pDL17 and pDL14) for dsDNA and oligonucleotide-mediated recombineering, respectively. The plasmid pDL14 contains a dominant negative mutS^K622A allele that suppresses methyl-directed mismatch repair (MMR). The coexpression of λRed and mutS^K622A provides efficient oligonucleotide-mediated recombineering in the presence of active host MMR. The expression of λRed was placed under the control of the tightly regulated P_rhaB promoter. Because of their low expression level under uninduced conditions, both plasmids could be maintained without elimination for multiple recombineering steps. The temperature-independent replication of the plasmids and control of λRed expression by l-rhamnose allow for all procedures to be performed at 37 °C. Thus, the new plasmids are robust, convenient, and versatile tools for Escherichia coli genome editing.

Discovery of recombinases enables genome mining of cryptic biosynthetic gene clusters in Burkholderiales species

Natural products biosynthesized by cryptic gene clusters represent a largely untapped source for drug discovery. However, mining of these products by promoter engineering is restricted by the lack of streamlined genetic tools, especially in nonmodel biosynthetic gene cluster (BGC)-rich bacteria. Here, we describe the discovery of a pair of bacteriophage recombinases and application of recombinase-assisted promoter engineering to rapidly identify and activate several cryptic biosynthetic gene clusters in two Burkholderiales strains that currently lack effective genetic tools. Construction of an efficient genome engineering platform in a natural product producer expedites mining of cryptic BGCs in their native backgrounds, and host melioration for yield or structure optimization. This strategy enables potentially scalable discovery of novel metabolites with intriguing bioactivities from many other bacteria.

Bacterial genomes encode numerous cryptic biosynthetic gene clusters (BGCs) that represent a largely untapped source of drugs or pesticides. Mining of the cryptic products is limited by the unavailability of streamlined genetic tools in native producers. Precise genome engineering using bacteriophage recombinases is particularly useful for genome mining. However, recombinases are usually host-specific. The genome-guided discovery of novel recombinases and their transient expression could boost cryptic BGC mining. Herein, we reported a genetic system employing Red recombinases from Burkholderiales strain DSM 7029 for efficient genome engineering in several Burkholderiales species that currently lack effective genetic tools. Using specialized recombinases-assisted in situ insertion of functional promoters, we successfully mined five cryptic nonribosomal peptide synthetase/polyketide synthase BGCs, two of which were silent. Two classes of lipopeptides, glidopeptins and rhizomides, were identified through extensive spectroscopic characterization. This recombinase expression strategy offers utility within other bacteria species, allowing bioprospecting for potentially scalable discovery of novel metabolites with attractive bioactivities.

An Engineered Virus Library as a Resource for the Spectrum-wide Exploration of Virus and Vector Diversity

Adenoviruses (Ads) are large human-pathogenic double-stranded DNA (dsDNA) viruses presenting an enormous natural diversity associated with a broad variety of diseases. However, only a small fraction of adenoviruses has been explored in basic virology and biomedical research, highlighting the need to develop robust and adaptable methodologies and resources. We developed a method for high-throughput direct cloning and engineering of adenoviral genomes from different sources utilizing advanced linear-linear homologous recombination (LLHR) and linear-circular homologous recombination (LCHR). We describe 34 cloned adenoviral genomes originating from clinical samples, which were characterized by next-generation sequencing (NGS). We anticipate that this recombineering strategy and the engineered adenovirus library will provide an approach to study basic and clinical virology. High-throughput screening (HTS) of the reporter-tagged Ad library in a panel of cell lines including osteosarcoma disease-specific cell lines revealed alternative virus types with enhanced transduction and oncolysis efficiencies. This highlights the usefulness of this resource.

Multiplex Genome Editing in Escherichia coli

Lambda Red recombineering is an easy and efficient method for generating genetic modifications in Escherichia coli. For gene deletions, lambda Red recombineering is combined with the use of selectable markers, which are removed through the action of, e.g., flippase (Flp) recombinase. This PCR-based engineering method has also been applied to a number of other bacteria. In this chapter, we describe a recently developed one plasmid-based method as well as the use of a strain with genomically integrated recombineering genes, which significantly speeds up the engineering of strains with multiple genomic alterations.

The contribution of homology arms to nuclease-assisted genome engineering

Designer nucleases like CRISPR/Cas9 enable fluent site-directed damage or small mutations in many genomes. Strategies for their use to achieve more complex tasks like regional exchanges for gene humanization or the establishment of conditional alleles are still emerging. To optimize Cas9-assisted targeting, we measured the relationship between targeting frequency and homology length in targeting constructs using a hypoxanthine-guanine phosphoribosyl-transferase assay in mouse embryonic stem cells. Targeting frequency with supercoiled plasmids improved steeply up to 2 kb total homology and continued to increase with even longer homology arms, thereby implying that Cas9-assisted targeting efficiencies can be improved using homology arms of 1 kb or greater. To humanize the Kmt2d gene, we built a hybrid mouse/human targeting construct in a bacterial artificial chromosome by recombineering. To simplify the possible outcomes, we employed a single Cas9 cleavage strategy and best achieved the intended 42 kb regional exchange with a targeting construct including a very long homology arm to recombine ∼42 kb away from the cleavage site. We recommend the use of long homology arm targeting constructs for accurate and efficient complex genome engineering, particularly when combined with the simplifying advantages of using just one Cas9 cleavage at the genome target site.

Direct and Inverted Repeat stimulated excision (DIRex): Simple, single-step, and scar-free mutagenesis of bacterial genes

The need for generating precisely designed mutations is common in genetics, biochemistry, and molecular biology. Here, I describe a new λ Red recombineering method (Direct and Inverted Repeat stimulated excision; DIRex) for fast and easy generation of single point mutations, small insertions or replacements as well as deletions of any size, in bacterial genes. The method does not leave any resistance marker or scar sequence and requires only one transformation to generate a semi-stable intermediate insertion mutant. Spontaneous excision of the intermediate efficiently and accurately generates the final mutant. In addition, the intermediate is transferable between strains by generalized transductions, enabling transfer of the mutation into multiple strains without repeating the recombineering step. Existing methods that can be used to accomplish similar results are either (i) more complicated to design, (ii) more limited in what mutation types can be made, or (iii) require expression of extrinsic factors in addition to λ Red. I demonstrate the utility of the method by generating several deletions, small insertions/replacements, and single nucleotide exchanges in Escherichia coli and Salmonella enterica. Furthermore, the design parameters that influence the excision frequency and the success rate of generating desired point mutations have been examined to determine design guidelines for optimal efficiency.

Duplication-Insertion Recombineering: a fast and scar-free method for efficient transfer of multiple mutations in bacteria

We have developed a new λ Red recombineering methodology for generating transient selection markers that can be used to transfer mutations between bacterial strains of both Escherichia coli and Salmonella enterica. The method is fast, simple and allows for the construction of strains with several mutations without any unwanted sequence changes (scar-free). The method uses λ Red recombineering to generate a marker-held tandem duplication, termed Duplication-Insertion (Dup-In). The Dup-Ins can easily be transferred between strains by generalized transduction and are subsequently rapidly lost by homologous recombination between the two copies of the duplicated sequence, leaving no scar sequence or antibiotic resistance cassette behind. We demonstrate the utility of the method by generating several Dup-Ins in E. coli and S. enterica to transfer genetically linked mutations in both essential and non-essential genes. We have successfully used this methodology to re-construct mutants found after various types of selections, and to introduce foreign genes into the two species. Furthermore, recombineering with two overlapping fragments was as efficient as recombineering with the corresponding single large fragment, allowing more complicated constructions without the need for overlap extension PCR.

Novel Technologies for Optimal Strain Breeding

The implementation of a knowledge-based bioeconomy requires the rapid development of highly efficient microbial production strains that are able to convert renewable carbon sources to value-added products, such as bulk and fine chemicals, pharmaceuticals, or proteins at industrial scale. Starting from classical strain breeding by random mutagenesis and screening in the 1950s via rational design by metabolic engineering initiated in the 1970s, a range of powerful new technologies have been developed in the past two decades that can revolutionize future strain engineering. In particular, next-generation sequencing technologies combined with new methods of genome engineering and high-throughput screening based on genetically encoded biosensors have allowed for new concepts. In this chapter, selected new technologies relevant for breeding microbial production strains with a special emphasis on amino acid producers will be summarized.

DNA annealing by Redβ is insufficient for homologous recombination and the additional requirements involve intra- and inter-molecular interactions

Single strand annealing proteins (SSAPs) like Redβ initiate homologous recombination by annealing complementary DNA strands. We show that C-terminally truncated Redβ, whilst still able to promote annealing and nucleoprotein filament formation, is unable to mediate homologous recombination. Mutations of the C-terminal domain were evaluated using both single- and double stranded (ss and ds) substrates in recombination assays. Mutations of critical amino acids affected either dsDNA recombination or both ssDNA and dsDNA recombination indicating two separable functions, one of which is critical for dsDNA recombination and the second for recombination per se. As evaluated by co-immunoprecipitation experiments, the dsDNA recombination function relates to the Redα-Redβ protein-protein interaction, which requires not only contacts in the C-terminal domain but also a region near the N-terminus. Because the nucleoprotein filament formed with C-terminally truncated Redβ has altered properties, the second C-terminal function could be due to an interaction required for functional filaments. Alternatively the second C-terminal function could indicate a requirement for a Redβ-host factor interaction. These data further advance the model for Red recombination and the proposition that Redβ and RAD52 SSAPs share ancestral and mechanistic roots.

Examining a DNA Replication Requirement for Bacteriophage λ Red- and Rac Prophage RecET-Promoted Recombination in Escherichia coli

Recombineering, in vivo genetic engineering with bacteriophage homologous recombination systems, is a powerful technique for making genetic modifications in bacteria. Two systems widely used in Escherichia coli are the Red system from phage λ and RecET from the defective Rac prophage. We investigated the in vivo dependence of recombineering on DNA replication of the recombining substrate using plasmid targets. For λ Red recombination, when DNA replication of a circular target plasmid is prevented, recombination with single-stranded DNA oligonucleotides is greatly reduced compared to that under replicating conditions. For RecET recombination, when DNA replication of the targeted plasmid is prevented, the recombination frequency is also reduced, to a level identical to that seen for the Red system in the absence of replication. The very low level of oligonucleotide recombination observed in the absence of any phage recombination functions is the same in the presence or absence of DNA replication. In contrast, both the Red and RecET systems recombine a nonreplicating linear dimer plasmid with high efficiency to yield a circular monomer. Therefore, the DNA replication requirement is substrate dependent. Our data are consistent with recombination by both the Red and RecET systems occurring predominately by single-strand annealing rather than by strand invasion.

IMPORTANCE

Bacteriophage homologous recombination systems are widely used for in vivo genetic engineering in bacteria. Single- or double-stranded linear DNA substrates containing short flanking homologies to chromosome targets are used to generate precise and accurate genetic modifications when introduced into bacteria expressing phage recombinases. Understanding the molecular mechanism of these recombination systems will facilitate improvements in the technology. Here, two phage-specific systems are shown to require exposure of complementary single-strand homologous targets for efficient recombination; these single-strand regions may be created during DNA replication or by single-strand exonuclease digestion of linear duplex DNA. Previously, in vitro studies reported that these recombinases promote the single-strand annealing of two complementary DNAs and also strand invasion of a single DNA strand into duplex DNA to create a three-stranded region. Here, in vivo experiments show that recombinase-mediated annealing of complementary single-stranded DNA is the predominant recombination pathway in E. coli.

Red-mediated recombineering of Salmonella enterica genomes

The mutagenesis of enterobacterial genomes using phage λ Red recombinase functions is a rapid and versatile experimental tool. In addition to the rapid generation of deletions in the genome of Salmonella enterica, variations of the method allow site-directed mutagenesis, generation of reporter fusions, generation of chimeric genes, or transplantation of regulatory elements directly in the chromosome. We describe the application of these approaches with focus on practical aspects and critical steps.

Molecular Cloning Designer Simulator (MCDS): All-in-one molecular cloning and genetic engineering design, simulation and management software for complex synthetic biology and metabolic engineering projects

Molecular Cloning Designer Simulator (MCDS) is a powerful new all-in-one cloning and genetic engineering design, simulation and management software platform developed for complex synthetic biology and metabolic engineering projects. In addition to standard functions, it has a number of features that are either unique, or are not found in combination in any one software package: (1) it has a novel interactive flow-chart user interface for complex multi-step processes, allowing an integrated overview of the whole project; (2) it can perform a user-defined workflow of cloning steps in a single execution of the software; (3) it can handle multiple types of genetic recombineering, a technique that is rapidly replacing classical cloning for many applications; (4) it includes experimental information to conveniently guide wet lab work; and (5) it can store results and comments to allow the tracking and management of the whole project in one platform. MCDS is freely available from https://mcds.codeplex.com.

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MCDS is an all-in-one in silico design, simulation and management platform.

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MCDS supports the design, simulation management of most cloning and recombineering technologies.

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MCDS has a novel interactive flowchart that allows simpler and more precise transactions.

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MCDS enables complete information integrity and consistency by keeping all details in one file.

λ Recombination and Recombineering

The bacteriophage λ Red homologous recombination system has been studied over the past 50 years as a model system to define the mechanistic details of how organisms exchange DNA segments that share extended regions of homology. The λ Red system proved useful as a system to study because recombinants could be easily generated by co-infection of genetically marked phages. What emerged from these studies was the recognition that replication of phage DNA was required for substantial Red-promoted recombination in vivo, and the critical role that double-stranded DNA ends play in allowing the Red proteins access to the phage DNA chromosomes. In the past 16 years, however, the λ Red recombination system has gained a new notoriety. When expressed independently of other λ functions, the Red system is able to promote recombination of linear DNA containing limited regions of homology (∼50 bp) with the Escherichia coli chromosome, a process known as recombineering. This review explains how the Red system works during a phage infection, and how it is utilized to make chromosomal modifications of E. coli with such efficiency that it changed the nature and number of genetic manipulations possible, leading to advances in bacterial genomics, metabolic engineering, and eukaryotic genetics.

Room temperature electrocompetent bacterial cells improve DNA transformation and recombineering efficiency

Bacterial competent cells are essential for cloning, construction of DNA libraries, and mutagenesis in every molecular biology laboratory. Among various transformation methods, electroporation is found to own the best transformation efficiency. Previous electroporation methods are based on washing and electroporating the bacterial cells in ice-cold condition that make them fragile and prone to death. Here we present simple temperature shift based methods that improve DNA transformation and recombineering efficiency in E. coli and several other gram-negative bacteria thereby economizing time and cost. Increased transformation efficiency of large DNA molecules is a significant advantage that might facilitate the cloning of large fragments from genomic DNA preparations and metagenomics samples.

REPLACR-mutagenesis, a one-step method for site-directed mutagenesis by recombineering

Mutagenesis is an important tool to study gene regulation, model disease-causing mutations and for functional characterisation of proteins. Most of the current methods for mutagenesis involve multiple step procedures. One of the most accurate methods for genetically altering DNA is recombineering, which uses bacteria expressing viral recombination proteins. Recently, the use of in vitro seamless assembly systems using purified enzymes for multiple-fragment cloning as well as mutagenesis is gaining ground. Although these in vitro isothermal reactions are useful when cloning multiple fragments, for site-directed mutagenesis it is unnecessary. Moreover, the use of purified enzymes in vitro is not only expensive but also more inaccurate than the high-fidelity recombination inside bacteria. Here we present a single-step method, named REPLACR-mutagenesis (Recombineering of Ends of linearised PLAsmids after PCR), for creating mutations (deletions, substitutions and additions) in plasmids by in vivo recombineering. REPLACR-mutagenesis only involves transformation of PCR products in bacteria expressing Red/ET recombineering proteins. Modifications in a variety of plasmids up to bacterial artificial chromosomes (BACs; 144 kb deletion) have been achieved by this method. The presented method is more robust, involves fewer steps and is cost-efficient.

Seven gene deletions in seven days: Fast generation of Escherichia coli strains tolerant to acetate and osmotic stress

Generation of multiple genomic alterations is currently a time consuming process. Here, a method was established that enables highly efficient and simultaneous deletion of multiple genes in Escherichia coli. A temperature sensitive plasmid containing arabinose inducible lambda Red recombineering genes and a rhamnose inducible flippase recombinase was constructed to facilitate fast marker-free deletions. To further speed up the procedure, we integrated the arabinose inducible lambda Red recombineering genes and the rhamnose inducible FLP into the genome of E. coli K-12 MG1655. This system enables growth at 37 °C, thereby facilitating removal of integrated antibiotic cassettes and deletion of additional genes in the same day. Phosphorothioated primers were demonstrated to enable simultaneous deletions during one round of electroporation. Utilizing these methods, we constructed strains in which four to seven genes were deleted in E. coli W and E. coli K-12. The growth rate of an E. coli K-12 quintuple deletion strain was significantly improved in the presence of high concentrations of acetate and NaCl. In conclusion, we have generated a method that enables efficient and simultaneous deletion of multiple genes in several E. coli variants. The method enables deletion of up to seven genes in as little as seven days.

Bacterial Recombineering: Genome Engineering via Phage-Based Homologous Recombination

The ability to specifically modify bacterial genomes in a precise and efficient manner is highly desired in various fields, ranging from molecular genetics to metabolic engineering and synthetic biology. Much has changed from the initial realization that phage-derived genes may be employed for such tasks to today, where recombineering enables complex genetic edits within a genome or a population. Here, we review the major developments leading to recombineering becoming the method of choice for in situ bacterial genome editing while highlighting the various applications of recombineering in pushing the boundaries of synthetic biology. We also present the current understanding of the mechanism of recombineering. Finally, we discuss in detail issues surrounding recombineering efficiency and future directions for recombineering-based genome editing.

The no-SCAR (Scarless Cas9 Assisted Recombineering) system for genome editing in Escherichia coli

Genome engineering methods in E. coli allow for easy to perform manipulations of the chromosome in vivo with the assistance of the λ-Red recombinase system. These methods generally rely on the insertion of an antibiotic resistance cassette followed by removal of the same cassette, resulting in a two-step procedure for genomic manipulations. Here we describe a method and plasmid system that can edit the genome of E. coli without chromosomal markers. This system, known as Scarless Cas9 Assisted Recombineering (no-SCAR), uses λ-Red to facilitate genomic integration of donor DNA and double stranded DNA cleavage by Cas9 to counterselect against wild-type cells. We show that point mutations, gene deletions, and short sequence insertions were efficiently performed in several genomic loci in a single-step with regards to the chromosome and did not leave behind scar sites. The single-guide RNA encoding plasmid can be easily cured due to its temperature sensitive origin of replication, allowing for iterative chromosomal manipulations of the same strain, as is often required in metabolic engineering. In addition, we demonstrate the ability to efficiently cure the second plasmid in the system by targeting with Cas9, leaving the cells plasmid-free.

A Single-Strand Annealing Protein Clamps DNA to Detect and Secure Homology

Repair of DNA breaks by single-strand annealing (SSA) is a major mechanism for the maintenance of genomic integrity. SSA is promoted by proteins (single-strand-annealing proteins [SSAPs]), such as eukaryotic RAD52 and λ phage Redβ. These proteins use a short single-stranded region to find sequence identity and initiate homologous recombination. However, it is unclear how SSAPs detect homology and catalyze annealing. Using single-molecule experiments, we provide evidence that homology is recognized by Redβ monomers that weakly hold single DNA strands together. Once annealing begins, dimerization of Redβ clamps the double-stranded region and nucleates nucleoprotein filament growth. In this manner, DNA clamping ensures and secures a successful detection for DNA sequence homology. The clamp is characterized by a structural change of Redβ and a remarkable stability against force up to 200 pN. Our findings not only present a detailed explanation for SSAP action but also identify the DNA clamp as a very stable, noncovalent, DNA-protein interaction.

Quantifying Impact of Chromosome Copy Number on Recombination in Escherichia coli

The ability to precisely and efficiently recombineer synthetic DNA into organisms of interest in a quantitative manner is a key requirement in genome engineering. Even though considerable effort has gone into the characterization of recombination in Escherichia coli, there is still substantial variability in reported recombination efficiencies. We hypothesized that this observed variability could, in part, be explained by the variability in chromosome copy number as well as the location of the replication forks relative to the recombination site. During rapid growth, E. coli cells may contain several pairs of open replication forks. While recombineered forks are resolving and segregating within the population, changes in apparent recombineering efficiency should be observed. In the case of dominant phenotypes, we predicted and then experimentally confirmed that the apparent recombination efficiency declined during recovery until complete segregation of recombineered and wild-type genomes had occurred. We observed the reverse trend for recessive phenotypes. The observed changes in apparent recombination efficiency were found to be in agreement with mathematical calculations based on our proposed mechanism. We also provide a model that can be used to estimate the total segregated recombination efficiency based on an initial efficiency and growth rate. These results emphasize the importance of employing quantitative strategies in the design of genome-scale engineering efforts.

Efficient conditional knockout targeting vector construction using co-selection BAC recombineering (CoSBR)

A simple and efficient strategy for Bacterial Artificial Chromosome (BAC) recombineering based on co-selection is described. We show that it is possible to efficiently modify two positions of a BAC simultaneously by co-transformation of a single-stranded DNA oligo and a double-stranded selection cassette. The use of co-selection BAC recombineering reduces the DNA manipulation needed to make a conditional knockout gene targeting vector to only two steps: a single round of BAC modification followed by a retrieval step.

Coupling the CRISPR/Cas9 System with Lambda Red Recombineering Enables Simplified Chromosomal Gene Replacement in Escherichia coli

To date, most genetic engineering approaches coupling the type II Streptococcus pyogenes clustered regularly interspaced short palindromic repeat (CRISPR)/Cas9 system to lambda Red recombineering have involved minor single nucleotide mutations. Here we show that procedures for carrying out more complex chromosomal gene replacements in Escherichia coli can be substantially enhanced through implementation of CRISPR/Cas9 genome editing. We developed a three-plasmid approach that allows not only highly efficient recombination of short single-stranded oligonucleotides but also replacement of multigene chromosomal stretches of DNA with large PCR products. By systematically challenging the proposed system with respect to the magnitude of chromosomal deletion and size of DNA insertion, we demonstrated DNA deletions of up to 19.4 kb, encompassing 19 nonessential chromosomal genes, and insertion of up to 3 kb of heterologous DNA with recombination efficiencies permitting mutant detection by colony PCR screening. Since CRISPR/Cas9-coupled recombineering does not rely on the use of chromosome-encoded antibiotic resistance, or flippase recombination for antibiotic marker recycling, our approach is simpler, less labor-intensive, and allows efficient production of gene replacement mutants that are both markerless and "scar"-less.

Differential requirements of singleplex and multiplex recombineering of large DNA constructs

Recombineering is an in vivo genetic engineering technique involving homologous recombination mediated by phage recombination proteins. The use of recombineering methodology is not limited by size and sequence constraints and therefore has enabled the streamlined construction of bacterial strains and multi-component plasmids. Recombineering applications commonly utilize singleplex strategies and the parameters are extensively tested. However, singleplex recombineering is not suitable for the modification of several loci in genome recoding and strain engineering exercises, which requires a multiplex recombineering design. Defining the main parameters affecting multiplex efficiency especially the insertion of multiple large genes is necessary to enable efficient large-scale modification of the genome. Here, we have tested different recombineering operational parameters of the lambda phage Red recombination system and compared singleplex and multiplex recombineering of large gene sized DNA cassettes. We have found that optimal multiplex recombination required long homology lengths in excess of 120 bp. However, efficient multiplexing was possible with only 60 bp of homology. Multiplex recombination was more limited by lower amounts of DNA than singleplex recombineering and was greatly enhanced by use of phosphorothioate protection of DNA. Exploring the mechanism of multiplexing revealed that efficient recombination required co-selection of an antibiotic marker and the presence of all three Red proteins. Building on these results, we substantially increased multiplex efficiency using an ExoVII deletion strain. Our findings elucidate key differences between singleplex and multiplex recombineering and provide important clues for further improving multiplex recombination efficiency.

HSV-I and the cellular DNA damage response

Peter Wildy first observed genetic recombination between strains of HSV in 1955. At the time, knowledge of DNA repair mechanisms was limited, and it has only been in the last decade that particular DNA damage response (DDR) pathways have been examined in the context of viral infections. One of the first reports addressing the interaction between a cellular DDR protein and HSV-1 was the observation by Lees-Miller et al. that DNA-dependent protein kinase catalytic subunit levels were depleted in an ICP0-dependent manner during Herpes simplex virus 1 infection. Since then, there have been numerous reports describing the interactions between HSV infection and cellular DDR pathways. Due to space limitations, this review will focus predominantly on the most recent observations regarding how HSV navigates a potentially hostile environment to replicate its genome.

Lambda red mediated gap repair utilizes a novel replicative intermediate in Escherichia coli

The lambda phage Red recombination system can mediate efficient homologous recombination in Escherichia coli, which is the basis of the DNA engineering technique termed recombineering. Red mediated insertion of DNA requires DNA replication, involves a single-stranded DNA intermediate and is more efficient on the lagging strand of the replication fork. Lagging strand recombination has also been postulated to explain the Red mediated repair of gapped plasmids by an Okazaki fragment gap filling model. Here, we demonstrate that gap repair involves a different strand independent mechanism. Gap repair assays examining the strand asymmetry of recombination did not show a lagging strand bias. Directly testing an ssDNA plasmid showed lagging strand recombination is possible but dsDNA plasmids did not employ this mechanism. Insertional recombination combined with gap repair also did not demonstrate preferential lagging strand bias, supporting a different gap repair mechanism. The predominant recombination route involved concerted insertion and subcloning though other routes also operated at lower frequencies. Simultaneous insertion of DNA resulted in modification of both strands and was unaffected by mutations to DNA polymerase I, responsible for Okazaki fragment maturation. The lower efficiency of an alternate Red mediated ends-in recombination pathway and the apparent lack of a Holliday junction intermediate suggested that gap repair does not involve a different Red recombination pathway. Our results may be explained by a novel replicative intermediate in gap repair that does not involve a replication fork. We exploited these observations by developing a new recombineering application based on concerted insertion and gap repair, termed SPI (subcloning plus insertion). SPI selected against empty vector background and selected for correct gap repair recombinants. We used SPI to simultaneously insert up to four different gene cassettes in a single recombineering reaction. Consequently, our findings have important implications for the understanding of E. coli replication and Red recombination.