Modified bases enable high-efficiency oligonucleotide-mediated allelic replacement via mismatch repair evasion

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.

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

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

IMPORTANCE

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

Strategies to identify and edit improvements in synthetic genome segments episomally

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

Advanced genome-editing technologies enable rapid and large-scale generation of genetic variants for strain engineering and synthetic biology

Targeted genome editing not only improves our understanding of fundamental rules in life sciences but also affords us versatile toolkits to improve industrially relevant phenotypes in various host cells. In this review, we summarize the recent endeavor to develop efficient genome-editing tools, and emphasize the utility of these tools to generate massive scale of genetic variants. We categorize these tools into traditional recombination-based tools, and more advanced CRISPR as well as RNA-based genome-editing tools. This diverse panel of sophisticated tools has been applied to accelerate strain engineering, upgrade biomanufacturing, and customize biosensing. In parallel with high-throughput phenotyping and AI-based optimization algorithms, we envision that genome-editing technologies will become a driving force to automate and streamline biological engineering, and empower us to address critical challenges in health, environment, energy, and sustainability.

Recombineering and MAGE

Recombination-mediated genetic engineering, also known as recombineering, is the genomic incorporation of homologous single-stranded or double-stranded DNA into bacterial genomes. Recombineering and its derivative methods have radically improved genome engineering capabilities, perhaps none more so than multiplex automated genome engineering (MAGE). MAGE is representative of a set of highly multiplexed single-stranded DNA-mediated technologies. First described in Escherichia coli, both MAGE and recombineering are being rapidly translated into diverse prokaryotes and even into eukaryotic cells. Together, this modern set of tools offers the promise of radically improving the scope and throughput of experimental biology by providing powerful new methods to ease the genetic manipulation of model and non-model organisms. In this Primer, we describe recombineering and MAGE, their optimal use, their diverse applications and methods for pairing them with other genetic editing tools. We then look forward to the future of genetic engineering.

Extensive trimming of short single-stranded DNA oligonucleotides during replication-coupled gene editing in mammalian cells

Through transfection of short single-stranded oligodeoxyribonucleotides (ssODNs) small genomic alterations can be introduced into mammalian cells with high precision. ssODNs integrate into the genome during DNA replication, but the resulting heteroduplex is prone to detection by DNA mismatch repair (MMR), which prevents effective gene modification. We have previously demonstrated that the suppressive action of MMR can be avoided when the mismatching nucleotide in the ssODN is a locked nucleic acid (LNA). Here, we reveal that LNA-modified ssODNs (LMOs) are not integrated as intact entities in mammalian cells, but are severely truncated before and after target hybridization. We found that single additional (non-LNA-modified) mutations in the 5’-arm of LMOs influenced targeting efficiencies negatively and activated the MMR pathway. In contrast, additional mutations in the 3’-arm did not affect targeting efficiencies and were not subject to MMR. Even more strikingly, homology in the 3’-arm was largely dispensable for effective targeting, suggestive for extensive 3’-end trimming. We propose a refined model for LMO-directed gene modification in mammalian cells that includes LMO degradation.

The first step of many gene editing approaches in mammalian cells is to generate a targeted DNA lesion. By administering a repair template as second step, endogenous DNA repair mechanisms can be misled to introduce specific gene variants. However, subtle gene modification can also be achieved with high precision through a one-action protocol in the absence of DNA breaks. We have shown before that short single-stranded DNA molecules (LMOs) are very useful to introduce and study genetic variants that may predispose patients to cancer. While LMOs are known to integrate into the genome during DNA replication, the precise mechanism is poorly understood. We targeted mouse embryonic stem cells with differently designed LMOs to examine their effectiveness and editing outcomes. Based on these results we conclude that the two LMO termini are processed at different moments during the gene editing process. While the 3’-arm is degraded prior to LMO binding to the target site, the 5’-arm is degraded afterwards. Counterintuitively we also observe that partial degradation of the 3’-arm increases targeting efficiencies. Taken together our data provides novel mechanistic insight into our understanding of replication-coupled gene editing and may guide future LMO design strategies.

Bacterial Genetic Engineering by Means of Recombineering for Reverse Genetics

Serving a robust platform for reverse genetics enabling the in vivo study of gene functions primarily in enterobacteriaceae, recombineering -or recombination-mediated genetic engineering-represents a powerful and relative straightforward genetic engineering tool. Catalyzed by components of bacteriophage-encoded homologous recombination systems and only requiring short ∼40–50 base homologies, the targeted and precise introduction of modifications (e.g., deletions, knockouts, insertions and point mutations) into the chromosome and other episomal replicons is empowered. Furthermore, by its ability to make use of both double- and single-stranded linear DNA editing substrates (e.g., PCR products or oligonucleotides, respectively), lengthy subcloning of specific DNA sequences is circumvented. Further, the more recent implementation of CRISPR-associated endonucleases has allowed for more efficient screening of successful recombinants by the selective purging of non-edited cells, as well as the creation of markerless and scarless mutants. In this review we discuss various recombineering strategies to promote different types of gene modifications, how they are best applied, and their possible pitfalls.

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.

A versatile platform strain for high-fidelity multiplex genome editing

Precision genome editing accelerates the discovery of the genetic determinants of phenotype and the engineering of novel behaviors in organisms. Advances in DNA synthesis and recombineering have enabled high-throughput engineering of genetic circuits and biosynthetic pathways via directed mutagenesis of bacterial chromosomes. However, the highest recombination efficiencies have to date been reported in persistent mutator strains, which suffer from reduced genomic fidelity. The absence of inducible transcriptional regulators in these strains also prevents concurrent control of genome engineering tools and engineered functions. Here, we introduce a new recombineering platform strain, BioDesignER, which incorporates (i) a refactored λ-Red recombination system that reduces toxicity and accelerates multi-cycle recombination, (ii) genetic modifications that boost recombination efficiency, and (iii) four independent inducible regulators to control engineered functions. These modifications resulted in single-cycle recombineering efficiencies of up to 25% with a 7-fold increase in recombineering fidelity compared to the widely used recombineering strain EcNR2. To facilitate genome engineering in BioDesignER, we have curated eight context--neutral genomic loci, termed Safe Sites, for stable gene expression and consistent recombination efficiency. BioDesignER is a platform to develop and optimize engineered cellular functions and can serve as a model to implement comparable recombination and regulatory systems in other bacteria.

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.

Directed evolution of multiple genomic loci allows the prediction of antibiotic resistance

Antibiotic development is frequently plagued by the rapid emergence of drug resistance. However, assessing the risk of resistance development in the preclinical stage is difficult. Standard laboratory evolution approaches explore only a small fraction of the sequence space and fail to identify exceedingly rare resistance mutations and combinations thereof. Therefore, new rapid and exhaustive methods are needed to accurately assess the potential of resistance evolution and uncover the underlying mutational mechanisms. Here, we introduce directed evolution with random genomic mutations (DIvERGE), a method that allows an up to million-fold increase in mutation rate along the full lengths of multiple predefined loci in a range of bacterial species. In a single day, DIvERGE generated specific mutation combinations, yielding clinically significant resistance against trimethoprim and ciprofloxacin. Many of these mutations have remained previously undetected or provide resistance in a species-specific manner. These results indicate pathogen-specific resistance mechanisms and the necessity of future narrow-spectrum antibacterial treatments. In contrast to prior claims, we detected the rapid emergence of resistance against gepotidacin, a novel antibiotic currently in clinical trials. Based on these properties, DIvERGE could be applicable to identify less resistance-prone antibiotics at an early stage of drug development. Finally, we discuss potential future applications of DIvERGE in synthetic and evolutionary biology.

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.

Orthogonal Ribosome Biofirewall

Biocontainment systems are crucial for preventing genetically modified organisms from escaping into natural ecosystems. Here, we describe the orthogonal ribosome biofirewall, which consists of an activation circuit and a degradation circuit. The activation circuit is a genetic AND gate based on activation of the encrypted pathway by the orthogonal ribosome in response to specific environmental signals. The degradation circuit is a genetic NOT gate with an output of I-SceI homing endonuclease, which conditionally degrades the orthogonal ribosome genes. We demonstrate that the activation circuit can be flexibly incorporated into genetic circuits and metabolic pathways for encryption. The plasmid-based encryption of the deoxychromoviridans pathway and the genome-based encryption of lacZ are tightly regulated and can decrease the expression to 7.3% and 7.8%, respectively. We validated the ability of the degradation circuit to decrease the expression levels of the target plasmids and the orthogonal rRNA (O-rRNA) plasmids to 0.8% in lab medium and 0.76% in nonsterile soil medium, respectively. Our orthogonal ribosome biofirewall is a versatile platform that can be useful in biosafety research and in the biotechnology industry.

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.

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.

CRISPR-based genome editing of clinically important Escherichia coli SE15 isolated from indwelling urinary catheters of patients

Urinary tract infections (UTIs) are clinically important problems that lead to serious morbidity and mortality, and indwelling urinary catheters are a major factor of UTIs. In this study, we applied clustered regularly interspaced short palindromic repeats (CRISPR) genome editing to generate ΔluxS mutant strains from clinical isolates of Escherichiacoli SE15, which is one of major pathogens and can cause colonization and biofilm formation in the catheter. A major regulatory pathway of such biofilm formation on medical devices is the quorum sensing mechanism via small molecule autoinducer-2 synthesized by LuxS enzyme. Here, we used the CRISPR-Cas9 system for precise deletion of luxS gene in clinical isolate E. coli SE15. To this end, we constructed a donor DNA for homologous recombination to delete 93 bases in the chromosomal target (luxS) and observed the success rate of luxS deletion to be 22.7 %. We conducted biofilm assay to observe decreased biofilm formation in the E. coil SE15 ΔluxS mutants compared to wild-type E. coil SE15. Quantitative real-time PCR analysis of E. coil SE15 ΔluxS mutants showed that the expression of luxS was below detection level. We also observed that the relative mRNA levels of biofilm-formation-related genes, such as mqsR, pgaBC and csgEF, were significantly decreased in E. coil SE15 ΔluxS mutants compared to wild-type. We conclude that genome editing by CRISPR-Cas9 system is an effective tool to dissect the molecular mechanism of biofilm formation in medically important strains, and the study may serve as a basis for developing novel medical intervention against UTIs caused by biofilm.

Leveraging modern DNA assembly techniques for rapid, markerless genome modification

The ability to alter the genomic material of a prokaryotic cell is necessary for experiments designed to define the biology of the organism. In addition, the production of biomolecules may be significantly improved by application of engineered prokaryotic host cells. Furthermore, in the age of synthetic biology, speed and efficiency are key factors when choosing a method for genome alteration. To address these needs, we have developed a method for modification of the Escherichia coli genome named FAST-GE for Fast Assembly-mediated Scarless Targeted Genome Editing. Traditional cloning steps such as plasmid transformation, propagation and isolation were eliminated. Instead, we developed a DNA assembly-based approach for generating scarless strain modifications, which may include point mutations, deletions and gene replacements, within 48 h after the receipt of polymerase chain reaction primers. The protocol uses established, but optimized, genome modification components such as I-SceI endonuclease to improve recombination efficiency and SacB as a counter-selection mechanism. All DNA-encoded components are assembled into a single allele-exchange vector named pDEL. We were able to rapidly modify the genomes of both E. coli B and K-12 strains with high efficiency. In principle, the method may be applied to other prokaryotic organisms capable of circular dsDNA uptake and homologous recombination.

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.

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.

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.

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.

Transient overexpression of DNA adenine methylase enables efficient and mobile genome engineering with reduced off-target effects

Homologous recombination of single-stranded oligonucleotides is a highly efficient process for introducing precise mutations into the genome of E. coli and other organisms when mismatch repair (MMR) is disabled. This can result in the rapid accumulation of off-target mutations that can mask desired phenotypes, especially when selections need to be employed following the generation of combinatorial libraries. While the use of inducible mutator phenotypes or other MMR evasion tactics have proven useful, reported methods either require non-mobile genetic modifications or costly oligonucleotides that also result in reduced efficiencies of replacement. Therefore a new system was developed, Transient Mutator Multiplex Automated Genome Engineering (TM-MAGE), that solves problems encountered in other methods for oligonucleotide-mediated recombination. TM-MAGE enables nearly equivalent efficiencies of allelic replacement to the use of strains with fully disabled MMR and with an approximately 12- to 33-fold lower off-target mutation rate. Furthermore, growth temperatures are not restricted and a version of the plasmid can be readily removed by sucrose counterselection. TM-MAGE was used to combinatorially reconstruct mutations found in evolved salt-tolerant strains, enabling the identification of causative mutations and isolation of strains with up to 75% increases in growth rate and greatly reduced lag times in 0.6 M NaCl.

Accelerated genome engineering through multiplexing

Throughout the biological sciences, the past 15 years have seen a push toward the analysis and engineering of biological systems at the organism level. Given the complexity of even the simplest organisms, though, to elicit a phenotype of interest often requires genotypic manipulation of several loci. By traditional means, sequential editing of genomic targets requires a significant investment of time and labor, as the desired editing event typically occurs at a very low frequency against an overwhelming unedited background. In recent years, the development of a suite of new techniques has greatly increased editing efficiency, opening up the possibility for multiple editing events to occur in parallel. Termed as multiplexed genome engineering, this approach to genome editing has greatly expanded the scope of possible genome manipulations in diverse hosts, ranging from bacteria to human cells. The enabling technologies for multiplexed genome engineering include oligonucleotide-based and nuclease-based methodologies, and their application has led to the great breadth of successful examples described in this review. While many technical challenges remain, there also exists a multiplicity of opportunities in this rapidly expanding field.

Genomes by design

Next-generation DNA sequencing has revealed the complete genome sequences of numerous organisms, establishing a fundamental and growing understanding of genetic variation and phenotypic diversity. Engineering at the gene, network and whole-genome scale aims to introduce targeted genetic changes both to explore emergent phenotypes and to introduce new functionalities. Expansion of these approaches into massively parallel platforms establishes the ability to generate targeted genome modifications, elucidating causal links between genotype and phenotype, as well as the ability to design and reprogramme organisms. In this Review, we explore techniques and applications in genome engineering, outlining key advances and defining challenges.

Superior Performance of Aptamer in Tumor Penetration over Antibody: Implication of Aptamer-Based Theranostics in Solid Tumors

Insufficient penetration of therapeutic agents into tumor tissues results in inadequate drug distribution and lower intracellular concentration of drugs, leading to the increase of drug resistance and resultant failure of cancer treatment. Targeted drug delivery to solid tumors followed by complete drug penetration and durable retention will significantly improve clinical outcomes of cancer therapy. Monoclonal antibodies have been commonly used in clinic for cancer treatment, but their limitation of penetrating into tumor tissues still remains because of their large size. Aptamers, as “chemical antibodies”, are 15-20 times smaller than antibodies. To explore whether aptamers are superior to antibodies in terms of tumor penetration, we carried out the first comprehensive study to compare the performance of an EpCAM aptamer with an EpCAM antibody in theranostic applications. Penetration and retention were studied in in vitro three-dimensional tumorspheres, in vivo live animal imaging and mouse colorectal cancer xenograft model. We found that the EpCAM aptamer can not only effectively penetrate into the tumorsphere cores but can also be retained by tumor sphere cells for at least 24 h, while limited tumor penetration by EpCAM antibody was observed after 4 h incubation. As observed from in vivo live animal imaging, EpCAM aptamers displayed a maximum tumor uptake at around 10 min followed by a rapid clearance after 80 min, while the signal of peak uptake and disappearance of antibody appeared at 3 h and 6 h after intravenous injection, respectively. The signal of PEGylated EpCAM aptamers in xenograft tumors was sustained for 26 h, which was 4.3-fold longer than that of the EpCAM antibody. Consistently, there were 1.67-fold and 6.6-fold higher accumulation of PEGylated aptamer in xenograft tumors than that of antibody, at 3 h and 24 h after intravenous administration, respectively. In addition, the aptamer achieved at least a 4-time better tumor penetration in xenograft tumors than that of the antibody at a 200 μm distances from the blood vessels 3 h after intravenous injection. Taken together, these data indicate that aptmers are superior to antibodies in cancer theranostics due to their better tumor penetration, more homogeneous distribution and longer retention in tumor sites. Thus, aptamers are promising agents for targeted tumor therapeutics and molecular imaging.

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.

Replicative DNA polymerase δ but not ε proofreads errors in Cis and in Trans

It is now well established that in yeast, and likely most eukaryotic organisms, initial DNA replication of the leading strand is by DNA polymerase ε and of the lagging strand by DNA polymerase δ. However, the role of Pol δ in replication of the leading strand is uncertain. In this work, we use a reporter system in Saccharomyces cerevisiae to measure mutation rates at specific base pairs in order to determine the effect of heterozygous or homozygous proofreading-defective mutants of either Pol ε or Pol δ in diploid strains. We find that wild-type Pol ε molecules cannot proofread errors created by proofreading-defective Pol ε molecules, whereas Pol δ can not only proofread errors created by proofreading-defective Pol δ molecules, but can also proofread errors created by Pol ε-defective molecules. These results suggest that any interruption in DNA synthesis on the leading strand is likely to result in completion by Pol δ and also explain the higher mutation rates observed in Pol δ-proofreading mutants compared to Pol ε-proofreading defective mutants. For strains reverting via AT→GC, TA→GC, CG→AT, and GC→AT mutations, we find in addition a strong effect of gene orientation on mutation rate in proofreading-defective strains and demonstrate that much of this orientation dependence is due to differential efficiencies of mispair elongation. We also find that a 3′-terminal 8 oxoG, unlike a 3′-terminal G, is efficiently extended opposite an A and is not subject to proofreading. Proofreading mutations have been shown to result in tumor formation in both mice and humans; the results presented here can help explain the properties exhibited by those proofreading mutants.

Many DNA polymerases are able to proofread their errors: after incorporation of a wrong base, the resulting mispair invokes an exonuclease activity of the polymerase that removes the mispaired base and allows replication to continue. Elimination of the proofreading activity thus results in much higher mutation rates. We demonstrate that the two major replicative DNA polymerases in yeast, Pol δ and Pol ε, have different proofreading abilities. In diploid cells, Pol ε is not able to proofread errors created by other Pol ε molecules, whereas Pol δ can proofread not only errors created by other Pol δ molecules but also errors created by Pol ε molecules. We also find that mispaired bases not corrected by proofreading have much different likelihoods of being extended, depending on the particular base-base mismatch. In humans, defects in Pol δ or Pol ε proofreading can lead to cancer, and these results help explain the formation of those tumors and the finding that Pol ε mutants seem to be found as frequently, or more so, in human tumors as Pol δ mutants.

A versatile and highly efficient method for scarless genome editing in Escherichia coli and Salmonella enterica

Background

Recently developed methods for genome editing in bacteria take advantage of the introduction of double-strand breaks by I-SceI in a mutation cassette to select for cells in which homologous recombination has healed the break and introduced a desired mutation. This elegantly designed method did not work well in our hands for most genes.

Results

We corrected a mutation in the gene encoding I-SceI that compromised the function of a previously used Red helper plasmid. Further, we found that transcription extending into the mutation cassette interferes with cleavage by I-SceI. Addition of two transcription terminators upstream of the cleavage site dramatically increases the efficiency of genome editing. We also developed an improved method for modification of essential genes. Inclusion of a segment of the essential gene consisting of synonymous codons restores an open reading frame when the mutation cassette is integrated into the genome and decreases the frequency of recombination events that fail to incorporate the desired mutation. The optimized protocol takes only 5 days and has been 100% successful for over 100 genomic modifications in our hands.

Conclusions

The method we describe here is reliable and versatile, enabling various types of genome editing in Escherichia coli and Salmonella enterica by straightforward modifications of the mutation cassette. We provide detailed descriptions of the methods as well as designs for insertions, deletions, and introduction of point mutations.

Vulnerabilities on the lagging-strand template: opportunities for mobile elements

Mobile genetic elements have the ability to move between positions in a genome. Some of these elements are capable of targeting one of the template strands during DNA replication. Examples found in bacteria include (a) Red recombination mediated by bacteriophage λ, (b) integration of group II mobile introns that reverse splice and reverse transcribe into DNA, (c) HUH endonuclease elements that move as single-stranded DNA, and (d) Tn7, a DNA cut-and-paste transposon that uses a target-site-selecting protein to target transposition into certain forms of DNA replication. In all of these examples, the lagging-strand template appears to be targeted using a variety of features specific to this strand. These features appear especially available in certain situations, such as when replication forks stall or collapse. In this review, we address the idea that features specific to the lagging-strand template represent vulnerabilities that are capitalized on by mobile genetic elements.

Precision genome engineering in lactic acid bacteria

Innovative new genome engineering technologies for manipulating chromosomes have appeared in the last decade. One of these technologies, recombination mediated genetic engineering (recombineering) allows for precision DNA engineering of chromosomes and plasmids in Escherichia coli. Single-stranded DNA recombineering (SSDR) allows for the generation of subtle mutations without the need for selection and without leaving behind any foreign DNA. In this review we discuss the application of SSDR technology in lactic acid bacteria, with an emphasis on key factors that were critical to move this technology from E. coli into Lactobacillus reuteri and Lactococcus lactis. We also provide a blueprint for how to proceed if one is attempting to establish SSDR technology in a lactic acid bacterium. The emergence of CRISPR-Cas technology in genome engineering and its potential application to enhancing SSDR in lactic acid bacteria is discussed. The ability to perform precision genome engineering in medically and industrially important lactic acid bacteria will allow for the genetic improvement of strains without compromising safety.

MODEST: a web-based design tool for oligonucleotide-mediated genome engineering and recombineering

Recombineering and multiplex automated genome engineering (MAGE) offer the possibility to rapidly modify multiple genomic or plasmid sites at high efficiencies. This enables efficient creation of genetic variants including both single mutants with specifically targeted modifications as well as combinatorial cell libraries. Manual design of oligonucleotides for these approaches can be tedious, time-consuming, and may not be practical for larger projects targeting many genomic sites. At present, the change from a desired phenotype (e.g. altered expression of a specific protein) to a designed MAGE oligo, which confers the corresponding genetic change, is performed manually. To address these challenges, we have developed the MAGE Oligo Design Tool (MODEST). This web-based tool allows designing of MAGE oligos for (i) tuning translation rates by modifying the ribosomal binding site, (ii) generating translational gene knockouts and (iii) introducing other coding or non-coding mutations, including amino acid substitutions, insertions, deletions and point mutations. The tool automatically designs oligos based on desired genotypic or phenotypic changes defined by the user, which can be used for high efficiency recombineering and MAGE. MODEST is available for free and is open to all users at http://modest.biosustain.dtu.dk.

A simple and effective method for construction of Escherichia coli strains proficient for genome engineering

Multiplex genome engineering is a standalone recombineering tool for large-scale programming and accelerated evolution of cells. However, this advanced genome engineering technique has been limited to use in selected bacterial strains. We developed a simple and effective strain-independent method for effective genome engineering in Escherichia coli. The method involves introducing a suicide plasmid carrying the λ Red recombination system into the mutS gene. The suicide plasmid can be excised from the chromosome via selection in the absence of antibiotics, thus allowing transient inactivation of the mismatch repair system during genome engineering. In addition, we developed another suicide plasmid that enables integration of large DNA fragments into the lacZ genomic locus. These features enable this system to be applied in the exploitation of the benefits of genome engineering in synthetic biology, as well as the metabolic engineering of different strains of E. coli.

Bacterial cellular engineering by genome editing and gene silencing

Genome editing is an important technology for bacterial cellular engineering, which is commonly conducted by homologous recombination-based procedures, including gene knockout (disruption), knock-in (insertion), and allelic exchange. In addition, some new recombination-independent approaches have emerged that utilize catalytic RNAs, artificial nucleases, nucleic acid analogs, and peptide nucleic acids. Apart from these methods, which directly modify the genomic structure, an alternative approach is to conditionally modify the gene expression profile at the posttranscriptional level without altering the genomes. This is performed by expressing antisense RNAs to knock down (silence) target mRNAs in vivo. This review describes the features and recent advances on methods used in genomic engineering and silencing technologies that are advantageously used for bacterial cellular engineering.

Conditional DNA repair mutants enable highly precise genome engineering

Oligonucleotide-mediated multiplex genome engineering is an important tool for bacterial genome editing. The efficient application of this technique requires the inactivation of the endogenous methyl-directed mismatch repair system that in turn leads to a drastically elevated genomic mutation rate and the consequent accumulation of undesired off-target mutations. Here, we present a novel strategy for mismatch repair evasion using temperature-sensitive DNA repair mutants and temporal inactivation of the mismatch repair protein complex in Escherichia coli. Our method relies on the transient suppression of DNA repair during mismatch carrying oligonucleotide integration. Using temperature-sensitive control of methyl-directed mismatch repair protein activity during multiplex genome engineering, we reduced the number of off-target mutations by 85%, concurrently maintaining highly efficient and unbiased allelic replacement.

Programmable removal of bacterial strains by use of genome-targeting CRISPR-Cas systems

CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems in bacteria and archaea employ CRISPR RNAs to specifically recognize the complementary DNA of foreign invaders, leading to sequence-specific cleavage or degradation of the target DNA. Recent work has shown that the accidental or intentional targeting of the bacterial genome is cytotoxic and can lead to cell death. Here, we have demonstrated that genome targeting with CRISPR-Cas systems can be employed for the sequence-specific and titratable removal of individual bacterial strains and species. Using the type I-E CRISPR-Cas system in Escherichia coli as a model, we found that this effect could be elicited using native or imported systems and was similarly potent regardless of the genomic location, strand, or transcriptional activity of the target sequence. Furthermore, the specificity of targeting with CRISPR RNAs could readily distinguish between even highly similar strains in pure or mixed cultures. Finally, varying the collection of delivered CRISPR RNAs could quantitatively control the relative number of individual strains within a mixed culture. Critically, the observed selectivity and programmability of bacterial removal would be virtually impossible with traditional antibiotics, bacteriophages, selectable markers, or tailored growth conditions. Once delivery challenges are addressed, we envision that this approach could offer a novel means to quantitatively control the composition of environmental and industrial microbial consortia and may open new avenues for the development of “smart” antibiotics that circumvent multidrug resistance and differentiate between pathogenic and beneficial microorganisms.

Controlling the composition of microbial populations is a critical aspect in medicine, biotechnology, and environmental cycles. While different antimicrobial strategies, such as antibiotics, antimicrobial peptides, and lytic bacteriophages, offer partial solutions, what remains elusive is a generalized and programmable strategy that can distinguish between even closely related microorganisms and that allows for fine control over the composition of a microbial population. This study demonstrates that RNA-directed immune systems in bacteria and archaea called CRISPR-Cas systems can provide such a strategy. These systems can be employed to selectively and quantitatively remove individual bacterial strains based purely on sequence information, creating opportunities in the treatment of multidrug-resistant infections, the control of industrial fermentations, and the study of microbial consortia.

Programmable removal of bacterial strains by use of genome-targeting CRISPR-Cas systems

CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems in bacteria and archaea employ CRISPR RNAs to specifically recognize the complementary DNA of foreign invaders, leading to sequence-specific cleavage or degradation of the target DNA. Recent work has shown that the accidental or intentional targeting of the bacterial genome is cytotoxic and can lead to cell death. Here, we have demonstrated that genome targeting with CRISPR-Cas systems can be employed for the sequence-specific and titratable removal of individual bacterial strains and species. Using the type I-E CRISPR-Cas system in Escherichia coli as a model, we found that this effect could be elicited using native or imported systems and was similarly potent regardless of the genomic location, strand, or transcriptional activity of the target sequence. Furthermore, the specificity of targeting with CRISPR RNAs could readily distinguish between even highly similar strains in pure or mixed cultures. Finally, varying the collection of delivered CRISPR RNAs could quantitatively control the relative number of individual strains within a mixed culture. Critically, the observed selectivity and programmability of bacterial removal would be virtually impossible with traditional antibiotics, bacteriophages, selectable markers, or tailored growth conditions. Once delivery challenges are addressed, we envision that this approach could offer a novel means to quantitatively control the composition of environmental and industrial microbial consortia and may open new avenues for the development of "smart" antibiotics that circumvent multidrug resistance and differentiate between pathogenic and beneficial microorganisms.

IMPORTANCE

Controlling the composition of microbial populations is a critical aspect in medicine, biotechnology, and environmental cycles. While different antimicrobial strategies, such as antibiotics, antimicrobial peptides, and lytic bacteriophages, offer partial solutions, what remains elusive is a generalized and programmable strategy that can distinguish between even closely related microorganisms and that allows for fine control over the composition of a microbial population. This study demonstrates that RNA-directed immune systems in bacteria and archaea called CRISPR-Cas systems can provide such a strategy. These systems can be employed to selectively and quantitatively remove individual bacterial strains based purely on sequence information, creating opportunities in the treatment of multidrug-resistant infections, the control of industrial fermentations, and the study of microbial consortia.

Rational optimization of tolC as a powerful dual selectable marker for genome engineering

Selection has been invaluable for genetic manipulation, although counter-selection has historically exhibited limited robustness and convenience. TolC, an outer membrane pore involved in transmembrane transport in E. coli, has been implemented as a selectable/counter-selectable marker, but counter-selection escape frequency using colicin E1 precludes using tolC for inefficient genetic manipulations and/or with large libraries. Here, we leveraged unbiased deep sequencing of 96 independent lineages exhibiting counter-selection escape to identify loss-of-function mutations, which offered mechanistic insight and guided strain engineering to reduce counter-selection escape frequency by ∼40-fold. We fundamentally improved the tolC counter-selection by supplementing a second agent, vancomycin, which reduces counter-selection escape by 425-fold, compared colicin E1 alone. Combining these improvements in a mismatch repair proficient strain reduced counter-selection escape frequency by 1.3E6-fold in total, making tolC counter-selection as effective as most selectable markers, and adding a valuable tool to the genome editing toolbox. These improvements permitted us to perform stable and continuous rounds of selection/counter-selection using tolC, enabling replacement of 10 alleles without requiring genotypic screening for the first time. Finally, we combined these advances to create an optimized E. coli strain for genome engineering that is ∼10-fold more efficient at achieving allelic diversity than previous best practices.

Yeast oligo-mediated genome engineering (YOGE)

High-frequency oligonucleotide-directed recombination engineering (recombineering) has enabled rapid modification of several prokaryotic genomes to date. Here, we present a method for oligonucleotide-mediated recombineering in the model eukaryote and industrial production host Saccharomyces cerevisiae , which we call yeast oligo-mediated genome engineering (YOGE). Through a combination of overexpression and knockouts of relevant genes and optimization of transformation and oligonucleotide designs, we achieve high gene-modification frequencies at levels that only require screening of dozens of cells. We demonstrate the robustness of our approach in three divergent yeast strains, including those involved in industrial production of biobased chemicals. Furthermore, YOGE can be iteratively executed via cycling to generate genomic libraries up to 10 (5) individuals at each round for diversity generation. YOGE cycling alone or in combination with phenotypic selections or endonuclease-based negative genotypic selections can be used to generate modified alleles easily in yeast populations with high frequencies.

Multiplex iterative plasmid engineering for combinatorial optimization of metabolic pathways and diversification of protein coding sequences

Engineering complex biological systems typically requires combinatorial optimization to achieve the desired functionality. Here, we present Multiplex Iterative Plasmid Engineering (MIPE), which is a highly efficient and customized method for combinatorial diversification of plasmid sequences. MIPE exploits ssDNA mediated λ Red recombineering for the introduction of mutations, allowing it to target several sites simultaneously and generate libraries of up to 10(7) sequences in one reaction. We also describe "restriction digestion mediated co-selection (RD CoS)", which enables MIPE to produce enhanced recombineering efficiencies with greatly simplified coselection procedures. To demonstrate this approach, we applied MIPE to fine-tune gene expression level in the 5-gene riboflavin biosynthetic pathway and successfully isolated a clone with 2.67-fold improved production in less than a week. We further demonstrated the ability of MIPE for highly multiplexed diversification of protein coding sequence by simultaneously targeting 23 codons scattered along the 750 bp sequence. We anticipate this method to benefit the optimization of diverse biological systems in synthetic biology and metabolic engineering.

Genome-scale engineering for systems and synthetic biology

This review provides an overview of methodologies and technologies enabling genome-scale engineering, focusing on the design, construction, and testing of modified genomes in a variety of organisms. Future applications for systems and synthetic biology are discussed.

Genome-modification technologies enable the rational engineering and perturbation of biological systems. Historically, these methods have been limited to gene insertions or mutations at random or at a few pre-defined locations across the genome. The handful of methods capable of targetedgene editing suffered from low efficiencies, significant labor costs, or both. Recent advances have dramatically expanded our ability to engineer cells in a directed and combinatorial manner. Here, we review current technologies and methodologies for genome-scale engineering, discuss the prospects for extending efficient genome modification to new hosts, and explore the implications of continued advances toward the development of flexibly programmable chasses, novel biochemistries, and safer organismal and ecological engineering.

Bacterial DNA polymerases participate in oligonucleotide recombination

Synthetic single-strand oligonucleotides (oligos) with homology to genomic DNA have proved to be highly effective for constructing designed mutations in targeted genomes, a process referred to as recombineering. The cellular functions important for this type of homologous recombination have yet to be determined. Towards this end, we have identified Escherichia coli functions that process the recombining oligo and affect bacteriophage λ Red-mediated oligo recombination. To determine the nature of oligo processing during recombination, each oligo contained multiple nucleotide changes: a single base change allowing recombinant selection, and silent changes serving as genetic markers to determine the extent of oligo processing during the recombination. Such oligos were often not incorporated into the host chromosome intact; many were partially degraded in the process of recombination. The position and number of these silent nucleotide changes within the oligo strongly affect both oligo processing and recombination frequency. Exonucleases, especially those associated with DNA Polymerases I and III, affect inheritance of the silent nucleotide changes in the oligos. We demonstrate for the first time that the major DNA polymerases (Pol I and Pol III) and DNA ligase are directly involved with oligo recombination.

AXM mutagenesis: an efficient means for the production of libraries for directed evolution of proteins

Affinity maturation is an important part of the recombinant antibody development process. There are several well-established approaches for generating libraries of mutated antibody genes for affinity maturation, but these approaches are generally too laborious or expensive to allow high-throughput, parallel processing of multiple antibodies. Here, we describe a scalable approach that enables the generation of libraries with greater than 10(8) clones from a single Escherichia coli transformation. In our method, a mutated DNA fragment is produced using PCR conditions that promote nucleotide misincorporation into newly synthesized DNA. In the PCR reaction, one of the primers contains at least three phosphorothioate linkages at its 5' end, and treatment of the PCR product with a 5' to 3' exonuclease is used to preferentially remove the strand synthesized with the non-modified primer, resulting in a single-stranded DNA fragment. This fragment then serves as a megaprimer to prime DNA synthesis on a uracilated, circular, single-stranded template in a Kunkel-like mutagenesis reaction that biases nucleotide base-changes between the megaprimer and uracilated DNA sequence in favor of the in vitro synthesized megaprimer. This method eliminates the inefficient subcloning steps that are normally required for the construction of affinity maturation libraries from randomly mutagenized antibody genes.

Genome-scale genetic engineering in Escherichia coli

Genome engineering has been developed to create useful strains for biological studies and industrial uses. However, a continuous challenge remained in the field: technical limitations in high-throughput screening and precise manipulation of strains. Today, technical improvements have made genome engineering more rapid and efficient. This review introduces recent advances in genome engineering technologies applied to Escherichia coli as well as multiplex automated genome engineering (MAGE), a recent technique proposed as a powerful toolkit due to its straightforward process, rapid experimental procedures, and highly efficient properties.