Enhanced levels of lambda Red-mediated recombinants in mismatch repair mutants

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

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

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

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

CRISPR/Cas9-Assisted Seamless Genome Editing in Lactobacillus plantarum and Its Application in N-Acetylglucosamine Production

Lactobacillus plantarum is a potential starter and health-promoting probiotic bacterium. Effective, precise, and diverse genome editing of Lactobacillus plantarum without introducing exogenous genes or plasmids is of great importance. In this study, CRISPR/Cas9-assisted double-stranded DNA (dsDNA) and single-stranded DNA (ssDNA) recombineering was established in L. plantarum WCFS1 to seamlessly edit the genome, including gene knockouts, insertions, and point mutations. To optimize our editing method, phosphorothioate modification was used to improve the dsDNA insertion, and adenine-specific methyltransferase was used to improve the ssDNA recombination efficiency. These strategies were applied to engineer L. plantarum WCFS1 toward producing N-acetylglucosamine (GlcNAc). nagB was truncated to eliminate the reverse reaction of fructose-6-phosphate (F6P) to glucosamine 6-phosphate (GlcN-6P). Riboswitch replacement and point mutation in glmS1 were introduced to relieve feedback repression. The resulting strain produced 797.3 mg/liter GlcNAc without introducing exogenous genes or plasmids. This strategy may contribute to the available methods for precise and diverse genetic engineering in lactic acid bacteria and boost strain engineering for more applications.IMPORTANCE CRISPR/Cas9-assisted recombineering is restricted in lactic acid bacteria because of the lack of available antibiotics and vectors. In this study, a seamless genome editing method was carried out in Lactobacillus plantarum using CRISPR/Cas9-assisted double-stranded DNA (dsDNA) and single-stranded DNA (ssDNA) recombineering, and recombination efficiency was effectively improved by endogenous adenine-specific methyltransferase overexpression. L. plantarum WCFS1 produced 797.3 mg/liter N-acetylglucosamine (GlcNAc) through reinforcement of the GlcNAc pathway, without introducing exogenous genes or plasmids. This seamless editing strategy, combined with the potential exogenous GlcNAc-producing pathway, makes this strain an attractive candidate for industrial use in the future.

Improved sgRNA design in bacteria via genome-wide activity profiling

CRISPR/Cas9 is a promising tool in prokaryotic genome engineering, but its success is limited by the widely varying on-target activity of single guide RNAs (sgRNAs). Based on the association of CRISPR/Cas9-induced DNA cleavage with cellular lethality, we systematically profiled sgRNA activity by co-expressing a genome-scale library (∼70 000 sgRNAs) with Cas9 or its specificity-improved mutant in Escherichia coli. Based on this large-scale dataset, we constructed a comprehensive and high-density sgRNA activity map, which enables selecting highly active sgRNAs for any locus across the genome in this model organism. We also identified ‘resistant’ genomic loci with respect to CRISPR/Cas9 activity, notwithstanding the highly accessible DNA in bacterial cells. Moreover, we found that previous sgRNA activity prediction models that were trained on mammalian cell datasets were inadequate when coping with our results, highlighting the key limitations and biases of previous models. We hence developed an integrated algorithm to accurately predict highly effective sgRNAs, aiming to facilitate CRISPR/Cas9-based genome engineering, screenings and antimicrobials design in bacteria. We also isolated the important sgRNA features that contribute to DNA cleavage and characterized their key differences among wild type Cas9 and its mutant, shedding light on the biophysical mechanisms of the CRISPR/Cas9 system.

Construction of a novel Escherichia coli expression system: relocation of lpxA from chromosome to a constitutive expression vector

The selective marker in the plasmid-based expression system is usually a gene that encodes an antibiotic-resistant protein; therefore, the antibiotic has to add to maintain the plasmid when growing the bacteria. This antibiotic addition would lead to increase of production cost and the environment contamination. In this study, a novel Escherichia coli expression system, the lpxA deletion mutant harboring an lpxA-carrying vector, was developed. To develop this system, three plasmids pCas9Cre, pTF-A-UD, and pRSFCmlpxA were constructed. The plasmid pCas9Cre produces enzymes Cas9, λ-Red, and Cre and can be cured by growing at 42 °C; pTF-A-UD contains several DNA fragments required for deleting the chromosomal lpxA and can be cured by adding isopropyl-D-thiogalactopyranoside; pRSFCmlpxA contains the lpxA mutant lpxA123 and CamR. When E. coli were transformed with these three plasmids, the chromosomal lpxA and the CamR in pRSFCmlpxA can be efficiently removed, resulting in an E. coli lpxA mutant harboring pRSFlpxA. The lpxA is essential for the growth of E. coli; its relocation from chromosome to a constitutive expression vector is an ideal strategy to maintain the vector without antibiotic addition. The lpxA123 in pRSFlpxA can complement the deletion of the chromosomal lpxA and provide a strong selective pressure to maintain the plasmid pRSFlpxA. This study provides an experimental evidence that this novel expression system is convenient and efficient to use and can be used to improve L-threonine biosynthesis in the wild type E. coli MG1655 and an L-threonine producing E. coli TWF006.

A Stress Response Monitoring Lipoprotein Trafficking to the Outer Membrane

The outer membrane built by Gram-negative bacteria such as Escherichia coli forms a barrier that prevents antibiotics from entering the cell, limiting clinical options at a time of prevalent antibiotic resistance. Stress responses ensure that barrier integrity is continuously maintained. We have identified the Cpx signal transduction system as a stress response that monitors the trafficking of lipid-anchored lipoproteins to the outer membrane. These lipoproteins are needed by every machine that builds the outer membrane. Cpx monitors just one lipoprotein, NlpE, to detect the efficiency of lipoprotein trafficking in the cell. NlpE and Cpx were previously shown to play a role in resistance to copper. We show that copper blocks lipoprotein trafficking, reconciling old and new observations. Copper is an important element in innate immunity against pathogens, and our findings suggest that NlpE and Cpx help E. coli survive the assault of copper on a key outer membrane assembly pathway.

Gram-negative bacteria produce lipid-anchored lipoproteins that are trafficked to their outer membrane (OM). These lipoproteins are essential components in each of the molecular machines that build the OM, including the Bam machine that assembles β-barrel proteins and the Lpt pathway that transports lipopolysaccharide. Stress responses are known to monitor Bam and Lpt function, yet no stress system has been found that oversees the fundamental process of lipoprotein trafficking. We used genetic and chemical biology approaches to induce several different lipoprotein trafficking stresses in Escherichia coli. Our results identified the Cpx two-component system as a stress response for monitoring trafficking. Cpx is activated by trafficking defects and is required to protect the cell against the consequence of the resulting stress. The OM-targeted lipoprotein NlpE acts as a sensor that allows Cpx to gauge trafficking efficiency. We reveal that NlpE signals to Cpx while it is transiting the inner membrane (IM) en route to the OM and that only a small highly conserved N-terminal domain is required for signaling. We propose that defective trafficking causes NlpE to accumulate in the IM, activating Cpx to mount a transcriptional response that protects cells. Furthermore, we reconcile this new role of NlpE in signaling trafficking defects with its previously proposed role in sensing copper (Cu) stress by demonstrating that Cu impairs acylation of lipoproteins and, consequently, their trafficking to the OM.

Synthetic biology for evolutionary engineering: from perturbation of genotype to acquisition of desired phenotype

With the increased attention on bio-based industry, demands for techniques that enable fast and effective strain improvement have been dramatically increased. Evolutionary engineering, which is less dependent on biological information, has been applied to strain improvement. Currently, synthetic biology has made great innovations in evolutionary engineering, particularly in the development of synthetic tools for phenotypic perturbation. Furthermore, discovering biological parts with regulatory roles and devising novel genetic circuits have promoted high-throughput screening and selection. In this review, we first briefly explain basics of synthetic biology tools for mutagenesis and screening of improved variants, and then describe how these strategies have been improved and applied to phenotypic engineering. Evolutionary engineering using advanced synthetic biology tools will enable further innovation in phenotypic engineering through the development of novel genetic parts and assembly into well-designed logic circuits that perform complex tasks.

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.

Simple and efficient genome recombineering using kil counter-selection in Escherichia coli

Seamless modification of the Escherichia coli genome using positive selection / negative selection is widely used in metabolic engineering and functional genome analysis. Some excellent negative selection systems have been reported, of which tetA-sacB and inducible toxins system are prominent. To expand the existing negative selection toolkit, we constructed a new negative selection marker system based on kil gene of lambda prophage. The selection stringency of kil was measured and compared with the most widely used counter-selection gene, sacB, at the lacI, ack, and dbpa loci using different E. coli strains. At all these loci of tested strains, the selection stringency of kil significantly exceeds that of sacB by 2- to 28-fold. When dsDNA fragments were employed for recombination, the efficiency for isolating the correct recombinant of kil was significantly higher than that of sacB. This new negative selection system does not require special media or extended incubation time. However, our system cannot be used in host strains containing temperature-sensitive kil gene. A Red system providing plasmid without kil gene is recommended for use together with our system. Our counter-selection system is expected to be an addition to the engineering arsenal of E. coli.

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.

A new recombineering system for precise genome-editing in Shewanella oneidensis strain MR-1 using single-stranded oligonucleotides

Shewanella oneidensis MR-1 is an invaluable host for the discovery and engineering of pathways important for bioremediation of toxic and radioactive metals and understanding extracellular electron transfer. However, genetic manipulation is challenging due to the lack of genetic tools. Previously, the only reliable method used for introducing DNA into Shewanella spp. at high efficiency was bacterial conjugation, enabling transposon mutagenesis and targeted knockouts using suicide vectors for gene disruptions. Here, we describe development of a robust and simple electroporation method in S. oneidensis that allows an efficiency of ~4.0 x 10⁶ transformants/µg DNA. High transformation efficiency is maintained when cells are frozen for long term storage. In addition, we report a new prophage-mediated genome engineering (recombineering) system using a λ Red Beta homolog from Shewanella sp. W3-18-1. By targeting two different chromosomal alleles, we demonstrate its application for precise genome editing using single strand DNA oligonucleotides and show that an efficiency of ~5% recombinants among total cells can be obtained. This is the first effective and simple strategy for recombination with markerless mutations in S. oneidensis. Continued development of this recombinant technology will advance high-throughput and genome modification efforts to engineer and investigate S. oneidensis and other environmental bacteria.

Retroelement-Based Genome Editing and Evolution

While several genome editing methods exist, few are suitable for the continuous evolution of targeted sequences.  Here we develop bacterial retroelements known as "retrons" for the dynamic,  in vivo editing and mutagenesis of targeted genes. We first optimized retrons' ability to introduce preprogrammed mutations, optimizing both their expression and the host machinery that interacts with them to increase the incorporation frequency of mutations 78-fold over rates previously reported in synthetic systems. The optimized system is capable of simultaneously overwriting 13 separate positions spanning  a 31-base length, and is for the first time shown to yield targeted deletions and insertions. To engineer retrons as a tool to introduce novel, unprogrammed mutations in specific targeted regions, we expressed them under a mutagenic T7 RNA polymerase. This coupled mutagenic T7 RNA polymerase-retron system enabled the evolution of diverse variants of environmentally selected antibiotic resistance genes, producing mutation rates in the targeted region 190-fold higher than  background cellular mutation rates, potentially enabling the dynamic, continuous self-evolution of selected phenotypes.

Expression Patterns, Genomic Conservation and Input Into Developmental Regulation of the GGDEF/EAL/HD-GYP Domain Proteins in Streptomyces

To proliferate, antibiotic-producing Streptomyces undergo a complex developmental transition from vegetative growth to the production of aerial hyphae and spores. This morphological switch is controlled by the signaling molecule cyclic bis-(3',5') di-guanosine-mono-phosphate (c-di-GMP) that binds to the master developmental regulator, BldD, leading to repression of key sporulation genes during vegetative growth. However, a systematical analysis of all the GGDEF/EAL/HD-GYP proteins that control c-di-GMP levels in Streptomyces is still lacking. Here, we have FLAG-tagged all 10 c-di-GMP turnover proteins in Streptomyces venezuelae and characterized their expression patterns throughout the life cycle, revealing that the diguanylate cyclase (DGC) CdgB and the phosphodiesterase (PDE) RmdB are the most abundant GGDEF/EAL proteins. Moreover, we have deleted all the genes coding for c-di-GMP turnover enzymes individually and analyzed morphogenesis of the mutants in macrocolonies. We show that the composite GGDEF-EAL protein CdgC is an active DGC and that deletion of the DGCs cdgB and cdgC enhance sporulation whereas deletion of the PDEs rmdA and rmdB delay development in S. venezuelae. By comparing the pan genome of 93 fully sequenced Streptomyces species we show that the DGCs CdgA, CdgB, and CdgC, and the PDE RmdB represent the most conserved c-di-GMP-signaling proteins in the genus Streptomyces.

Optimization of Artificial Curcumin Biosynthesis in E. coli by Randomized 5'-UTR Sequences To Control the Multienzyme Pathway

One of the optimization strategies of an artificial biosynthetic metabolic flux with a multienzyme pathway is when the enzyme concentrations are present at the appropriate ratios rather than at their maximum expression. Thus, many recent research efforts have focused on the development of tools that fine-tune the enzyme expression, and these research efforts have facilitated the search for the optimum balance between pathway expression and cell viability. However, the rational approach has some limitations in finding the most optimized expression ratio in in vivo systems. In our study, we focused on fine-tuning the expression level of a six-enzyme reaction for the artificial biosynthesis of curcumin by screening a library of 5'-untranslational region (UTR) sequence mutants made by a multiplex automatic genome engineering (MAGE) tool. From the screening results, a variant (6M08rv) showed about a 38.2-fold improvement in the production of curcumin compared to the parent strain, in which the calculated expression levels of 4-coumarate:CoA ligase (4CL) and phenyldiketide-CoA synthase (DCS), two of the six enzymes, were much lower than those of the parent strain.

Targeted Nucleotide Editing Technologies for Microbial Metabolic Engineering

Since the emergence of programmable RNA-guided nucleases based on clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein (Cas) systems, genome editing technologies have become a simplified and versatile tool for genome editing in various organisms and cell types. Although genome editing enables efficient genome manipulations, such as gene disruptions, gene knockins, and chromosomal translocations via DNA double-strand break (DSB) repair in eukaryotes, DSBs induced by the CRISPR/Cas system are lethal or severely toxic to many microorganisms. Therefore, in many prokaryotes, including industrially useful microbes, the CRISPR/Cas system is often used as a negative selection component in combination with recombineering or other related strategies. Novel and revolutionary technologies have been recently developed to re-write targeted nucleotides (C:G to T:A and A:T to G:C) without DSBs and donor DNA templates. These technologies rely on the recruitment of deaminases at specific target loci using the nuclease-deficient CRISPR/Cas system. Here, the authors review and compare CRISPR-based genome editing, current base editing platforms and their spectra. The authors discuss how these technologies can be applied in various aspects of microbial metabolic engineering to overcome barriers to cellular regulation in prokaryotes.

CRISPR-guided DNA polymerases enable diversification of all nucleotides in a tunable window

The capacity to diversify genetic codes advances our ability to understand and engineer biological systems^1,2. A method for continuously diversifying user-defined regions of a genome would enable forward genetic approaches in systems that are not amenable to efficient homology-directed oligonucleotide integration. It would also facilitate the rapid evolution of biotechnologically useful phenotypes through accelerated and parallelized rounds of mutagenesis and selection, as well as cell-lineage tracking through barcode mutagenesis. Here we present EvolvR, a system that can continuously diversify all nucleotides within a tunable window length at user-defined loci. This is achieved by directly generating mutations using engineered DNA polymerases targeted to loci via CRISPR-guided nickases. We identified nickase and polymerase variants that offer a range of targeted mutation rates that are up to 7,770,000-fold greater than rates seen in wild-type cells, and editing windows with lengths of up to 350 nucleotides. We used EvolvR to identify novel ribosomal mutations that confer resistance to the antibiotic spectinomycin. Our results demonstrate that CRISPR-guided DNA polymerases enable multiplexed and continuous diversification of user-defined genomic loci, which will be useful for a broad range of basic and biotechnological applications.

CRISPR/Cas9 Assisted Multiplex Genome Editing Technique in Escherichia coli

Genome editing for site-specific chromosome modification is one of the most significant techniques in biological research. While conventional techniques usually deal with one genomic locus at a time, multiple genomic targets are often required to be modified to develop microbial cell factories. Thus, it is necessary to develop techniques for simultaneous editing of multiple loci. In this work, the authors develop a CRISPR/Cas9 assisted multiplex genome editing (CMGE) technique in Escherichia coli. With this editing method, all functional parts are assembled into replicable plasmids, and stringent inducible expression systems are used to control Cas9 gene expression, which is to decouple transformation from editing process to increase editing efficiency. A modular assembly strategy is designed to enable construction of the complex multi-gRNA plasmid. With this technique, two and three loci are able to be modified with 100% and 88.3% efficiencies, while four loci can be edited with more than 30%, which are the best results reported. Although developed in model organism, the strategy of CMGE can be adapted to other prokaryotic cells. This is a well designed and illustrated technique with no special requirement, can be used by any biological lab easily.

Absence of Curli in Soil-Persistent Escherichia coli Is Mediated by a C-di-GMP Signaling Defect and Suggests Evidence of Biofilm-Independent Niche Specialization

Escherichia coli is commonly viewed as a gastrointestinal commensal or pathogen although an increasing body of evidence suggests that it can persist in non-host environments as well. Curli are a major component of biofilm in many enteric bacteria including E. coli and are important for adherence to different biotic and abiotic surfaces. In this study we investigated curli production in a unique collection of soil-persistent E. coli isolates and examined the role of curli formation in environmental persistence. Although most soil-persistent E. coli were curli-positive, 10% of isolates were curli-negative (17 out of 170). Curli-producing E. coli (COB583, COB585, and BW25113) displayed significantly more attachment to quartz sand than the curli-negative strains. Long-term soil survival experiments indicated that curli production was not required for long-term survival in live soil (over 110 days), as a curli-negative mutant BW25113ΔcsgB had similar survival compared to wild type BW25113. Mutations in two genes associated with c-di-GMP metabolism, dgcE and pdeR, correlated with loss of curli in eight soil-persistent strains, although this did not significantly impair their survival in soil compared to curli-positive strains. Overall, the data indicate that curli-deficient and biofilm-defective strains, that also have a defect in attachment to quartz sand, are able to reside in soil for long periods of time thus pointing to the possibility that niches may exist in the soil that can support long-term survival independently of biofilm formation.

Endonuclease-independent DNA mismatch repair processes on the lagging strand

DNA mismatch repair (MMR) pathways coordinate the excision and re-synthesis of newly-replicated DNA if a mismatched base-pair has been identified by protein MutS or MutS homologues (MSHs) after replication. DNA excision during MMR is initiated at single-strand breaks (SSBs) in vitro, and several redundant processes have been observed in reconstituted systems which either require a pre-formed SSB in the DNA or require a mismatch-activated nicking endonuclease to introduce a SSB in order to initiate MMR. However, the conditions under which each of these processes may actually occur in living cells have remained obscured by the limitations of current MMR assays. Here we use a novel assay involving chemically-modified oligonucleotide probes to insert targeted DNA 'mismatches' directly into the genome of living bacteria to interrogate their replication-coupled repair processes quantitatively in a strand-, orientation-, and mismatched nucleotide-specific manner. This 'semi-protected oligonucleotide recombination' (SPORE) assay reveals direct evidence in Escherichia coli of an efficient endonuclease-independent MMR process on the lagging strand-a mechanism that has long-since been considered for lagging-strand repair but never directly shown until now. We find endonuclease-independent MMR is coordinated asymmetrically with respect to the replicating DNA-directed primarily from 3'- of the mismatch-and that repair coordinated from 3'- of the mismatch is in fact the primary mechanism of lagging-strand MMR. While further work is required to explore and identify the molecular requirements for this alternative endonuclease-independent MMR pathway, these findings made possible using the SPORE assay are the first direct report of this long-suspected mechanism in vivo.

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.

Sak4 of Phage HK620 Is a RecA Remote Homolog With Single-Strand Annealing Activity Stimulated by Its Cognate SSB Protein

Bacteriophages are remarkable for the wide diversity of proteins they encode to perform DNA replication and homologous recombination. Looking back at these ancestral forms of life may help understanding how similar proteins work in more sophisticated organisms. For instance, the Sak4 family is composed of proteins similar to the archaeal RadB protein, a Rad51 paralog. We have previously shown that Sak4 allowed single-strand annealing in vivo, but only weakly compared to the phage λ Redβ protein, highlighting putatively that Sak4 requires partners to be efficient. Here, we report that the purified Sak4 of phage HK620 infecting Escherichia coli is a poorly efficient annealase on its own. A distant homolog of SSB, which gene is usually next to the sak4 gene in various species of phages, highly stimulates its recombineering activity in vivo. In vitro, Sak4 binds single-stranded DNA and performs single-strand annealing in an ATP-dependent way. Remarkably, the single-strand annealing activity of Sak4 is stimulated by its cognate SSB. The last six C-terminal amino acids of this SSB are essential for the binding of Sak4 to SSB-covered single-stranded DNA, as well as for the stimulation of its annealase activity. Finally, expression of sak4 and ssb from HK620 can promote low-level of recombination in vivo, though Sak4 and its SSB are unable to promote strand exchange in vitro. Regarding its homology with RecA, Sak4 could represent a link between two previously distinct types of recombinases, i.e., annealases that help strand exchange proteins and strand exchange proteins themselves.

Deaminase-mediated multiplex genome editing in Escherichia coli

In eukaryotes, the CRISPR-Cas9 system has now been widely used as a revolutionary genome engineering tool1, 2. However, in prokaryotes, the use of nuclease-mediated genome editing tools has been limited to negative selection for the already modified cells because of its lethality3, 4. Here, we report on deaminase-mediated targeted nucleotide editing (Target-AID) 5 adopted in Escherichia coli. Cytidine deaminase PmCDA1 fused to the nuclease-deficient CRISPR-Cas9 system achieved specific point mutagenesis at the target sites in E. coli by introducing cytosine mutations without compromising cell growth. The cytosine-to-thymine substitutions were induced mainly within an approximately five-base window of target sequences on the protospacer adjacent motif-distal side, which can be shifted depending on the length of the single guide RNA sequence. Use of a uracil DNA glycosylase inhibitor 6 in combination with a degradation tag (LVA tag) 7 resulted in a robustly high mutation efficiency, which allowed simultaneous multiplex editing of six different genes. The major multi-copy transposase genes that consist of at least 41 loci were also simultaneously edited by using four target sequences. As this system does not rely on any additional or host-dependent factors, it may be readily applicable to a wide range of bacteria.

Efficient and Scalable Precision Genome Editing in Staphylococcus aureus through Conditional Recombineering and CRISPR/Cas9-Mediated Counterselection

Staphylococcus aureus is an important human pathogen, but studies of the organism have suffered from the lack of a robust tool set for its genetic and genomic manipulation. Here we report the development of a system for the facile and high-throughput genomic engineering of S. aureus using single-stranded DNA (ssDNA) oligonucleotide recombineering coupled with clustered regularly interspaced short palindromic repeat (CRISPR)/Cas9-mediated counterselection. We identify recombinase EF2132, derived from Enterococcus faecalis, as being capable of integrating single-stranded DNA oligonucleotides into the S. aureus genome. We found that EF2132 can readily mediate recombineering across multiple characterized strains (3 of 3 tested) and primary clinical isolates (6 of 6 tested), typically yielding thousands of recombinants per transformation. Surprisingly, we also found that some S. aureus strains are naturally recombinogenic at measurable frequencies when oligonucleotides are introduced by electroporation, even without exogenous recombinase expression. We construct a temperature-sensitive, two-vector system which enables conditional recombineering and CRISPR/Cas9-mediated counterselection in S. aureus without permanently introducing exogenous genetic material or unintended genetic lesions. We demonstrate the ability of this system to efficiently and precisely engineer point mutations and large single-gene deletions in the S. aureus genome and to yield highly enriched populations of engineered recombinants even in the absence of an externally selectable phenotype. By virtue of utilizing inexpensive, commercially synthesized synthetic DNA oligonucleotides as substrates for recombineering and counterselection, this system provides a scalable, versatile, precise, inexpensive, and generally useful tool for producing isogenic strains in S. aureus which will enable the high-throughput functional assessment of genome variation and gene function across multiple strain backgrounds.

Engineering genetic changes in bacteria is critical to understanding the function of particular genes or mutations but is currently a laborious and technically challenging process to perform for the important human pathogen Staphylococcus aureus. In an effort to develop methods which are rapid, easy, scalable, versatile, and inexpensive, here we describe a system for incorporating synthetic, mutagenic DNA molecules into the S. aureus genome and for eliminating cells that lack the engineered mutation. This method allows efficient, precise, and high-throughput genetic engineering of S. aureus strains and will facilitate studies seeking to address a variety of issues about the function of particular genes and specific mutations.

Precise Editing at DNA Replication Forks Enables Multiplex Genome Engineering in Eukaryotes

We describe a multiplex genome engineering technology in Saccharomyces cerevisiae based on annealing synthetic oligonucleotides at the lagging strand of DNA replication. The mechanism is independent of Rad51-directed homologous recombination and avoids the creation of double-strand DNA breaks, enabling precise chromosome modifications at single base-pair resolution with an efficiency of >40%, without unintended mutagenic changes at the targeted genetic loci. We observed the simultaneous incorporation of up to 12 oligonucleotides with as many as 60 targeted mutations in one transformation. Iterative transformations of a complex pool of oligonucleotides rapidly produced large combinatorial genomic diversity >10⁵. This method was used to diversify a heterologous β-carotene biosynthetic pathway that produced genetic variants with precise mutations in promoters, genes, and terminators, leading to altered carotenoid levels. Our approach of engineering the conserved processes of DNA replication, repair, and recombination could be automated and establishes a general strategy for multiplex combinatorial genome engineering in eukaryotes.

The Streptomyces master regulator BldD binds c-di-GMP sequentially to create a functional BldD2-(c-di-GMP)4 complex

Streptomyces are ubiquitous soil bacteria that undergo a complex developmental transition coinciding with their production of antibiotics. This transition is controlled by binding of a novel tetrameric form of the second messenger, 3΄-5΄ cyclic diguanylic acid (c-di-GMP) to the master repressor, BldD. In all domains of life, nucleotide-based second messengers allow a rapid integration of external and internal signals into regulatory pathways that control cellular responses to changing conditions. c-di-GMP can assume alternative oligomeric states to effect different functions, binding to effector proteins as monomers, intercalated dimers or, uniquely in the case of BldD, as a tetramer. However, at physiological concentrations c-di-GMP is a monomer and little is known about how higher oligomeric complexes assemble on effector proteins and if intermediates in assembly pathways have regulatory significance. Here, we show that c-di-GMP binds BldD using an ordered, sequential mechanism and that BldD function necessitates the assembly of the BldD₂-(c-di-GMP)₄ complex.

Systems Metabolic Engineering of Escherichia coli

Systems metabolic engineering, which recently emerged as metabolic engineering integrated with systems biology, synthetic biology, and evolutionary engineering, allows engineering of microorganisms on a systemic level for the production of valuable chemicals far beyond its native capabilities. Here, we review the strategies for systems metabolic engineering and particularly its applications in Escherichia coli. First, we cover the various tools developed for genetic manipulation in E. coli to increase the production titers of desired chemicals. Next, we detail the strategies for systems metabolic engineering in E. coli, covering the engineering of the native metabolism, the expansion of metabolism with synthetic pathways, and the process engineering aspects undertaken to achieve higher production titers of desired chemicals. Finally, we examine a couple of notable products as case studies produced in E. coli strains developed by systems metabolic engineering. The large portfolio of chemical products successfully produced by engineered E. coli listed here demonstrates the sheer capacity of what can be envisioned and achieved with respect to microbial production of chemicals. Systems metabolic engineering is no longer in its infancy; it is now widely employed and is also positioned to further embrace next-generation interdisciplinary principles and innovation for its upgrade. Systems metabolic engineering will play increasingly important roles in developing industrial strains including E. coli that are capable of efficiently producing natural and nonnatural chemicals and materials from renewable nonfood biomass.

A standardized workflow for surveying recombinases expands bacterial genome-editing capabilities

Bacterial recombineering typically relies on genomic incorporation of synthetic oligonucleotides as mediated by Escherichia coli λ phage recombinase β - an occurrence largely limited to enterobacterial strains. While a handful of similar recombinases have been documented, recombineering efficiencies usually fall short of expectations for practical use. In this work, we aimed to find an efficient Recβ homologue demonstrating activity in model soil bacterium Pseudomonas putida EM42. To this end, a genus-wide protein survey was conducted to identify putative recombinase candidates for study. Selected novel proteins were assayed in a standardized test to reveal their ability to introduce the K43T substitution into the rpsL gene of P. putida. An ERF superfamily protein, here termed Rec2, exhibited activity eightfold greater than that of the previous leading recombinase. To bolster these results, we demonstrated Rec2 ability to enter a range of mutations into the pyrF gene of P. putida at similar frequencies. Our results not only confirm the utility of Rec2 as a Recβ functional analogue within the P. putida model system, but also set a complete workflow for deploying recombineering in other bacterial strains/species. Implications range from genome editing of P. putida for metabolic engineering to extended applications within other Pseudomonads - and beyond.

A novel model to characterize structure and function of BRCA1

BRCA1 plays a central role in DNA repair. Although N-terminal RING and C-terminal BRCT domains are studied well, the functions of the central region of BRCA1 are poorly characterized. Here, we report a structural and functional analysis of BRCA1 alleles and functional human BRCA1 in chicken B-lymphocyte cell line DT40. The combination of "homologous recombineering" and "RT-cassette" enables modifications of chicken BRCA1 gene in Escherichia coli. Mutant BRCA1 knock-in DT40 cell lines were generated using BRCA1 mutation constructs by homologous recombination with a targeting efficiency of up to 100%. Our study demonstrated that deletion of motifs 2-9 BRCA1^{Δ/Δ181-1415} (Caenorhabditis elegans BRCA1 mimic) or deletion of motif 1 BRCA1^Δ/Δ126-136 decreased cell viability following cisplatin treatment. Furthermore, deletion of motifs 5 and 6 BRCA1^Δ/Δ525-881 within DNA-binding region, even the conserved 7-amino acid deletion BRCA1^Δ/Δ872-878 within motif 6, caused a decreased cell viability upon cisplatin treatment. Surprisingly, human BRCA1 is functional in DT40 cells as indicated by DNA damage-induced Rad 51 foci formation in human BRCA1 knock-in DT40 cells. These results demonstrate that those conserved motifs within the central region are essential for DNA repair functions of BRCA1. These findings provide a valuable tool for the development of new therapeutic modalities of breast cancer linked to BRCA1.

Efficient engineering of chromosomal ribosome binding site libraries in mismatch repair proficient Escherichia coli

Multiplexed gene expression optimization via modulation of gene translation efficiency through ribosome binding site (RBS) engineering is a valuable approach for optimizing artificial properties in bacteria, ranging from genetic circuits to production pathways. Established algorithms design smart RBS-libraries based on a single partially-degenerate sequence that efficiently samples the entire space of translation initiation rates. However, the sequence space that is accessible when integrating the library by CRISPR/Cas9-based genome editing is severely restricted by DNA mismatch repair (MMR) systems. MMR efficiency depends on the type and length of the mismatch and thus effectively removes potential library members from the pool. Rather than working in MMR-deficient strains, which accumulate off-target mutations, or depending on temporary MMR inactivation, which requires additional steps, we eliminate this limitation by developing a pre-selection rule of genome-library-optimized-sequences (GLOS) that enables introducing large functional diversity into MMR-proficient strains with sequences that are no longer subject to MMR-processing. We implement several GLOS-libraries in Escherichia coli and show that GLOS-libraries indeed retain diversity during genome editing and that such libraries can be used in complex genome editing operations such as concomitant deletions. We argue that this approach allows for stable and efficient fine tuning of chromosomal functions with minimal effort.

A 'Semi-Protected Oligonucleotide Recombination' Assay for DNA Mismatch Repair in vivo Suggests Different Modes of Repair for Lagging Strand Mismatches

In Escherichia coli, a DNA mismatch repair (MMR) pathway corrects errors that occur during DNA replication by coordinating the excision and re-synthesis of a long tract of the newly-replicated DNA between an epigenetic signal (a hemi-methylated d(GATC) site or a single-stranded nick) and the replication error after the error is identified by protein MutS. Recent observations suggest that this 'long-patch repair' between these sites is coordinated in the same direction of replication by the replisome. Here, we have developed a new assay that uniquely allows us to introduce targeted 'mismatches' directly into the replication fork via oligonucleotide recombination, examine the directionality of MMR, and quantify the nucleotide-dependence, sequence context-dependence, and strand-dependence of their repair in vivo-something otherwise nearly impossible to achieve. We find that repair of genomic lagging strand mismatches occurs bi-directionally in E. coli and that, while all MutS-recognized mismatches had been thought to be repaired in a consistent manner, the directional bias of repair and the effects of mutations in MutS are dependent on the molecular species of the mismatch. Because oligonucleotide recombination is routinely performed in both prokaryotic and eukaryotic cells, we expect this assay will be broadly applicable for investigating mechanisms of MMR in vivo.

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.

Rewriting the Genetic Code

The genetic code-the language used by cells to translate their genomes into proteins that perform many cellular functions-is highly conserved throughout natural life. Rewriting the genetic code could lead to new biological functions such as expanding protein chemistries with noncanonical amino acids (ncAAs) and genetically isolating synthetic organisms from natural organisms and viruses. It has long been possible to transiently produce proteins bearing ncAAs, but stabilizing an expanded genetic code for sustained function in vivo requires an integrated approach: creating recoded genomes and introducing new translation machinery that function together without compromising viability or clashing with endogenous pathways. In this review, we discuss design considerations and technologies for expanding the genetic code. The knowledge obtained by rewriting the genetic code will deepen our understanding of how genomes are designed and how the canonical genetic code evolved.

Translational Control of the SigR-Directed Oxidative Stress Response in Streptomyces via IF3-Mediated Repression of a Noncanonical GTC Start Codon

The major oxidative stress response in Streptomyces is controlled by the sigma factor SigR and its cognate antisigma factor RsrA, and SigR activity is tightly controlled through multiple mechanisms at both the transcriptional and posttranslational levels. Here we show that sigR has a highly unusual GTC start codon and that this leads to another level of SigR regulation, in which SigR translation is repressed by translation initiation factor 3 (IF3). Changing the GTC to a canonical start codon causes SigR to be overproduced relative to RsrA, resulting in unregulated and constitutive expression of the SigR regulon. Similarly, introducing IF3* mutations that impair its ability to repress SigR translation has the same effect. Thus, the noncanonical GTC sigR start codon and its repression by IF3 are critical for the correct and proper functioning of the oxidative stress regulatory system. sigR and rsrA are cotranscribed and translationally coupled, and it had therefore been assumed that SigR and RsrA are produced in stoichiometric amounts. Here we show that RsrA can be transcribed and translated independently of SigR, present evidence that RsrA is normally produced in excess of SigR, and describe the factors that determine SigR-RsrA stoichiometry.

In all sigma factor-antisigma factor regulatory switches, the relative abundance of the two proteins is critical to the proper functioning of the system. Many sigma-antisigma operons are cotranscribed and translationally coupled, leading to a generic assumption that the sigma and antisigma factors are produced in a fixed 1:1 ratio. In the case of sigR-rsrA, we show instead that the antisigma factor is produced in excess over the sigma factor, providing a buffer to prevent spurious release of sigma activity. This excess arises in part because sigR has an extremely rare noncanonical GTC start codon, and as a result, SigR translation initiation is repressed by IF3. This finding highlights the potential significance of noncanonical start codons, very few of which have been characterized experimentally. It also emphasizes the limitations of predicting start codons using bioinformatic approaches, which rely heavily on the assumption that ATG, GTG, and TTG are the only permissible start codons.

Optimizing complex phenotypes through model-guided multiplex genome engineering

We present a method for identifying genomic modifications that optimize a complex phenotype through multiplex genome engineering and predictive modeling. We apply our method to identify six single nucleotide mutations that recover 59% of the fitness defect exhibited by the 63-codon E. coli strain C321.∆A. By introducing targeted combinations of changes in multiplex we generate rich genotypic and phenotypic diversity and characterize clones using whole-genome sequencing and doubling time measurements. Regularized multivariate linear regression accurately quantifies individual allelic effects and overcomes bias from hitchhiking mutations and context-dependence of genome editing efficiency that would confound other strategies.

Scarless deletion of up to seven methyl-accepting chemotaxis genes with an optimized method highlights key function of CheM in Salmonella Typhimurium

Site-directed scarless mutagenesis is an essential tool of modern pathogenesis research. We describe an optimized two-step protocol for genome editing in Salmonella enterica serovar Typhimurium to enable multiple sequential mutagenesis steps in a single strain. The system is based on the λ Red recombinase-catalyzed integration of a selectable antibiotics resistance marker followed by replacement of this cassette. Markerless mutants are selected by expressing the meganuclease I-SceI which induces double-strand breaks in bacteria still harboring the resistance locus. Our new dual-functional plasmid pWRG730 allows for heat-inducible expression of the λ Red recombinase and tet-inducible production of I-SceI. Methyl-accepting chemotaxis proteins (MCP) are transmembrane chemoreceptors for a vast set of environmental signals including amino acids, sugars, ions and oxygen. Based on the sensory input of MCPs, chemotaxis is a key component for Salmonella virulence. To determine the contribution of individual MCPs we sequentially deleted seven MCP genes. The individual mutations were validated by PCR and genetic integrity of the final seven MCP mutant WRG279 was confirmed by whole genome sequencing. The successive MCP mutants were functionally tested in a HeLa cell infection model which revealed increased invasion rates for non-chemotactic mutants and strains lacking the MCP CheM (Tar). The phenotype of WRG279 was reversed with plasmid-based expression of CheM. The complemented WRG279 mutant showed also partially restored chemotaxis in swarming assays on semi-solid agar. Our optimized scarless deletion protocol enables efficient and precise manipulation of the Salmonella genome. As demonstrated with whole genome sequencing, multiple subsequent mutagenesis steps can be realized without the introduction of unwanted mutations. The sequential deletion of seven MCP genes revealed a significant role of CheM for the interaction of S. Typhimurium with host cells which might give new insights into mechanisms of Salmonella host cell sensing.

Automated electrotransformation of Escherichia coli on a digital microfluidic platform using bioactivated magnetic beads

This paper reports on the use of a digital microfluidic platform to perform multiplex automated genetic engineering (MAGE) cycles on droplets containing Escherichia coli cells. Bioactivated magnetic beads were employed for cell binding, washing, and media exchange in the preparation of electrocompetent cells in the electrowetting-on-dieletric (EWoD) platform. On-cartridge electroporation was used to deliver oligonucleotides into the cells. In addition to the optimization of a magnetic bead-based benchtop protocol for generating and transforming electrocompetent E. coli cells, we report on the implementation of this protocol in a fully automated digital microfluidic platform. Bead-based media exchange and electroporation pulse conditions were optimized on benchtop for transformation frequency to provide initial parameters for microfluidic device trials. Benchtop experiments comparing electrotransformation of free and bead-bound cells are presented. Our results suggest that dielectric shielding intrinsic to bead-bound cells significantly reduces electroporation field exposure efficiency. However, high transformation frequency can be maintained in the presence of magnetic beads through the application of more intense electroporation pulses. As a proof of concept, MAGE cycles were successfully performed on a commercial EWoD cartridge using variations of the optimal magnetic bead-based preparation procedure and pulse conditions determined by the benchtop results. Transformation frequencies up to 22% were achieved on benchtop; this frequency was matched within 1% (21%) by MAGE cycles on the microfluidic device. However, typical frequencies on the device remain lower, averaging 9% with a standard deviation of 9%. The presented results demonstrate the potential of digital microfluidics to perform complex and automated genetic engineering protocols.

Toward Genome-Based Metabolic Engineering in Bacteria

Prokaryotes modified stably on the genome are of great importance for production of fine and commodity chemicals. Traditional methods for genome engineering have long suffered from imprecision and low efficiencies, making construction of suitable high-producer strains laborious. Here, we review the recent advances in discovery and refinement of molecular precision engineering tools for genome-based metabolic engineering in bacteria for chemical production, with focus on the λ-Red recombineering and the clustered regularly interspaced short palindromic repeats/Cas9 nuclease systems. In conjunction, they enable the integration of in vitro-synthesized DNA segments into specified locations on the chromosome and allow for enrichment of rare mutants by elimination of unmodified wild-type cells. Combination with concurrently developing improvements in important accessory technologies such as DNA synthesis, high-throughput screening methods, regulatory element design, and metabolic pathway optimization tools has resulted in novel efficient microbial producer strains and given access to new metabolic products. These new tools have made and will likely continue to make a big impact on the bioengineering strategies that transform the chemical industry.

Genome-wide mapping of mutations at single-nucleotide resolution for protein, metabolic and genome engineering

Improvements in DNA synthesis and sequencing have underpinned comprehensive assessment of gene function in bacteria and eukaryotes. Genome-wide analyses require high-throughput methods to generate mutations and analyze their phenotypes, but approaches to date have been unable to efficiently link the effects of mutations in coding regions or promoter elements in a highly parallel fashion. We report that CRISPR-Cas9 gene editing in combination with massively parallel oligomer synthesis can enable trackable editing on a genome-wide scale. Our method, CRISPR-enabled trackable genome engineering (CREATE), links each guide RNA to homologous repair cassettes that both edit loci and function as barcodes to track genotype-phenotype relationships. We apply CREATE to site saturation mutagenesis for protein engineering, reconstruction of adaptive laboratory evolution experiments, and identification of stress tolerance and antibiotic resistance genes in bacteria. We provide preliminary evidence that CREATE will work in yeast. We also provide a webtool to design multiplex CREATE libraries.

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.

System-level genome editing in microbes

The release of the first complete microbial genome sequences at the end of the past century opened the way for functional genomics and systems-biology to uncover the genetic basis of various phenotypes. The surge of available sequence data facilitated the development of novel genome editing techniques for system-level analytical studies. Recombineering allowed unprecedented throughput and efficiency in microbial genome editing and the recent discovery and widespread use of RNA-guided endonucleases offered several further perspectives: (i) previously recalcitrant species became editable, (ii) the efficiency of recombineering could be elevated, and as a result (iii) diverse genomic libraries could be generated more effectively. Supporting recombineering by RNA-guided endonucleases has led to success stories in metabolic engineering, but their use for system-level analysis is mostly unexplored. For the full exploitation of opportunities that are offered by the genome editing proficiency, future development of large scale analytical procedures is also vitally needed.

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.

The Ssr protein (T1E_1405) from Pseudomonas putida DOT-T1E enables oligonucleotide-based recombineering in platform strain P. putida EM42

Some strains of the soil bacterium Pseudomonas putida have become in recent years platforms of choice for hosting biotransformations of industrial interest. Despite availability of many genetic tools for this microorganism, genomic editing of the cell factory P. putida EM42 (a derivative of reference strain KT2440) is still a time-consuming endeavor. In this work we have investigated the in vivo activity of the Ssr protein encoded by the open reading frame T1E_1405 from Pseudomonas putida DOT-T1E, a plausible functional homologue of the β protein of the Red recombination system of λ phage of Escherichia coli. A test based on the phenotypes of pyrF mutants of P. putida (the yeast's URA3 ortholog) was developed for quantifying the ability of Ssr to promote invasion of the genomic DNA replication fork by synthetic oligonucleotides. The efficiency of the process was measured by monitoring the inheritance of the changes entered into pyrF by oligonucleotides bearing mutated sequences. Ssr fostered short and long genomic deletions/insertions at considerable frequencies as well as single-base swaps not affected by mismatch repair. These results not only demonstrate the feasibility of recombineering in P. putida, but they also enable a suite of multiplexed genomic manipulations in this biotechnologically important bacterium.

A DNA Binding Protein Is Required for Viral Replication and Transcription in Bombyx mori Nucleopolyhedrovirus

A DNA-binding protein (DBP) [GenBank accession number: M63416] of Bombyx mori nuclear polyhedrosis virus (BmNPV) has been reported to be a regulatory factor in BmNPV, but its detailed functions remain unknown. In order to study the regulatory mechanism of DBP on viral proliferation, genome replication, and gene transcription, a BmNPV dbp gene knockout virus dbp-ko-Bacmid was generated by the means of Red recombination system. In addition, dbp-repaired virus dbp-re-Bacmid was constructed by the means of the Bac to Bac system. Then, the Bacmids were transfected into BmN cells. The results of this viral titer experiment revealed that the TCID50 of the dbp-ko-Bacmid was 0; however, the dbp-re-Bacmid was similar to the wtBacmid (p>0.05), indicating that the dbp-deficient would lead to failure in the assembly of virus particles. In the next step, Real-Time PCR was used to analyze the transcriptional phases of dbp gene in BmN cells, which had been infected with BmNPV. The results of the latter experiment revealed that the transcript of dbp gene was first detected at 3 h post-infection. Furthermore, the replication level of virus genome and the transcriptional level of virus early, late, and very late genes in BmN cells, which had been transfected with 3 kinds of Bacmids, were analyzed by Real-Time PCR. The demonstrating that the replication level of genome was lower than that of wtBacmid and dbp-re-Bacmid (p<0.01). The transcriptional level of dbp-ko-Bacmid early gene lef-3, ie-1, dnapol, late gene vp39 and very late gene p10 were statistically significantly lower than dbp-re-Bacmid and wtBacmid (p<0.01). The results presented are based on Western blot analysis, which indicated that the lack of dbp gene would lead to low expressions of lef3, vp39, and p10. In conclusion, dbp was not only essential for early viral replication, but also a viral gene that has a significant impact on transcription and expression during all periods of baculovirus life cycle.

Merlin: Computer-Aided Oligonucleotide Design for Large Scale Genome Engineering with MAGE

Genome engineering technologies now enable precise manipulation of organism genotype, but can be limited in scalability by their design requirements. Here we describe Merlin ( http://merlincad.org ), an open-source web-based tool to assist biologists in designing experiments using multiplex automated genome engineering (MAGE). Merlin provides methods to generate pools of single-stranded DNA oligonucleotides (oligos) for MAGE experiments by performing free energy calculation and BLAST scoring on a sliding window spanning the targeted site. These oligos are designed not only to improve recombination efficiency, but also to minimize off-target interactions. The application further assists experiment planning by reporting predicted allelic replacement rates after multiple MAGE cycles, and enables rapid result validation by generating primer sequences for multiplexed allele-specific colony PCR. Here we describe the Merlin oligo and primer design procedures and validate their functionality compared to OptMAGE by eliminating seven AvrII restriction sites from the Escherichia coli genome.

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.

LNA modification of single-stranded DNA oligonucleotides allows subtle gene modification in mismatch-repair-proficient cells

Synthetic single-stranded DNA oligonucleotides (ssODNs) can be used to generate subtle genetic modifications in eukaryotic and prokaryotic cells without the requirement for prior generation of DNA double-stranded breaks. However, DNA mismatch repair (MMR) suppresses the efficiency of gene modification by >100-fold. Here we present a commercially available ssODN design that evades MMR and enables subtle gene modification in MMR-proficient cells. The presence of locked nucleic acids (LNAs) in the ssODNs at mismatching bases, or also at directly adjacent bases, allowed 1-, 2-, or 3-bp substitutions in MMR-proficient mouse embryonic stem cells as effectively as in MMR-deficient cells. Additionally, in MMR-proficient Escherichia coli, LNA modification of the ssODNs enabled effective single-base-pair substitution. In vitro, LNA modification of mismatches precluded binding of purified E. coli MMR protein MutS. These findings make ssODN-directed gene modification particularly well suited for applications that require the evaluation of a large number of sequence variants with an easy selectable phenotype.

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

Currently available tools for multiplex bacterial genome engineering are optimized for a few laboratory model strains, demand extensive prior modification of the host strain, and lead to the accumulation of numerous off-target modifications. Building on prior development of multiplex automated genome engineering (MAGE), our work addresses these problems in a single framework. Using a dominant-negative mutant protein of the methyl-directed mismatch repair (MMR) system, we achieved a transient suppression of DNA repair in Escherichia coli, which is necessary for efficient oligonucleotide integration. By integrating all necessary components into a broad-host vector, we developed a new workflow we term pORTMAGE. It allows efficient modification of multiple loci, without any observable off-target mutagenesis and prior modification of the host genome. Because of the conserved nature of the bacterial MMR system, pORTMAGE simultaneously allows genome editing and mutant library generation in other biotechnologically and clinically relevant bacterial species. Finally, we applied pORTMAGE to study a set of antibiotic resistance-conferring mutations in Salmonella enterica and E. coli. Despite over 100 million y of divergence between the two species, mutational effects remained generally conserved. In sum, a single transformation of a pORTMAGE plasmid allows bacterial species of interest to become an efficient host for genome engineering. These advances pave the way toward biotechnological and therapeutic applications. Finally, pORTMAGE allows systematic comparison of mutational effects and epistasis across a wide range of bacterial species.