High-yield production of FK228 and new derivatives in a Burkholderia chassis.

FK228 (romidepsin) is the only natural histone deacetylases (HDACs) inhibitor approved by FDA to treat cutaneous and peripheral T-cell lymphoma. However, the limited supply and severe cardiotoxicity of FK228 underscore the importance to develop an effective synthetic biology platform for the manufacturing and fine-tuning of this drug lead. In this work, we constructed a Burkholderia chassis for the high-yield production of FK228-family (unnatural) natural products. By virtue of the optimized Burkholderia-specific recombineering system, the biosynthetic gene cluster (BGC) encoding the FK228-like skeleton thailandepsins (tdp) in Burkholderia thailandensis E264 was replaced with an attB integration site to afford the basal chassis KOGC1. The tdp BGC directly captured from E264 was hybridized with the FK228-encoding BGC (dep) using the versatile Red/ET technology. The hybrid BGC (tdp-dep) was integrated into the attB site of KOGC1, resulting in the heterologous expression of FK228. Remarkably, the titer reached 581 mg/L, which is 30-fold higher than that of native producer Chromobacterium violaceum No. 968. This success encouraged us to further engineer the NRPS modules 4 or 6 of hybrid tdp-dep BGC by domain units swapping strategy, and eight new FK228 derivatives (1-8) varying in the composition of amino acids were generated. Especially, the titers of 2 and 3 in KOGC1 were up to 985 mg/L and 453 mg/L, respectively. 2 and 3 displayed stronger cytotoxic activity than FK228. All in all, this work established a robust platform to produce FK228 and its new derivatives in sufficient quantities for anticancer drug development.

Improved terephthalic acid production from p-xylene using metabolically engineered Pseudomonas putida.

Terephthalic acid (TPA) is an important commodity chemical used as a monomer of polyethylene terephthalate (PET). Since a large quantity of PET is routinely manufactured and consumed worldwide, the development of sustainable biomanufacturing processes for its monomers (i.e. TPA and ethylene glycol) has recently gained much attention. In a previous study, we reported the development of a metabolically engineered Escherichia coli strain producing 6.7 g/L of TPA from p-xylene (pX) with a productivity and molar conversion yield of 0.278 g/L/h and 96.7 mol%, respectively. Here, we report metabolic engineering of Pseudomonas putida KT2440, a microbial chassis particularly suitable for the synthesis of aromatic compounds, for improved biocatalytic conversion of pX to TPA. To develop a plasmid-free, antibiotic-free, and inducer-free biocatalytic process for cost-competitive TPA production, all heterologous genes required for the synthetic pX-to-TPA bioconversion pathway were integrated into the chromosome of P. putida KT2440 by RecET-based markerless recombineering and overexpressed under the control of constitutive promoters. Next, TPA production was enhanced by integrating multiple copies of the heterologous genes to the ribosomal RNA genes through iteration of recombineering-based random integration and subsequent screening of high-performance strains. Finally, fed-batch fermentation process was optimized to further improve the performance of the engineered P. putida strain. As a result, 38.25 ± 0.11 g/L of TPA was produced from pX with a molar conversion yield of 99.6 ± 0.6%, which is equivalent to conversion of 99.3 ± 0.8 g pX to 154.6 ± 0.5 g TPA. This superior pX-to-TPA biotransformation process based on the engineered P. putida strain will pave the way to the commercial biomanufacturing of TPA in an industrial scale.

Characterization of a novel Escherichia coli recombineering selection/counterselection cassette.

Recombineering is a highly efficient DNA cloning and modification technique by using the recombinase-mediated homologous recombination. Selection/counterselection cassette is often used in chromosomal DNA or large episomal DNA manipulation, in which the selection marker is used for the first step cassette selection while deleting the target gene via allelic exchange, and the counterselection marker is used for the second step replacement of the cassette by the foreign DNA fragment. A variety of selection/counterselection cassettes are reported, however, the cassettes suffer from the shortcomings of the requirement of pre-engineered strain or specific culture medium. Herein, we report a novel S-tetR- PtetA-ccdB-aacC1-S selection/counterselection cassette that sidesteps the disadvantages. As a proof-of-concept, one-step gene cloning (0.7, 1.7, and 4.2 kb) and two-step Escherichia coli chromosomal gene knock-in (0.7 and 4.2 kb) were performed. The gene cloning and gene knock-in efficiencies are high up to 90%. The novel selection/counterselection cassette adds a powerful tool to the recombineering repertoire.

CRISPR/Cas9-assisted ssDNA recombineering for site-directed mutagenesis and saturation mutagenesis of plasmid-encoded genes.

Site-directed and saturation mutagenesis are critical DNA methodologies for studying protein structure and function. For plasmid-based gene mutation, PCR and overlap-extension PCR involve tedious cloning steps. When the plasmid size is large, PCR yield may be too low for cloning; and for saturation mutagenesis of a single codon, one experiment may not enough to generate all twenty coding variants. Oligo-mediated recombineering sidesteps the complicated cloning process by homologous recombination between a mutagenic oligo and its target site. However, the low recombineering efficiency and inability to select for the recombinant makes it necessary to screen a large number of clones. Herein, we describe two plasmid-based mutagenic strategies: CRISPR/Cas9-assisted ssDNA recombineering for site-directed mutagenesis (CRM) and saturation mutagenesis (CRSM). CRM and CRSM involve co-electroporation of target plasmid, sgRNA expression plasmid and mutagenic oligonucleotide into Escherichia coli cells with induced expression of λ-Red recombinase and Cas9, followed by plasmid extraction and characterization. We established CRM and CRSM via ampicillin resistance gene repair and mutagenesis of N-acetyl­D­neuraminic acid aldolase. The mutational efficiency was between 80 and 100% and all twenty amino acid coding variants were obtained at a target site via a single CRSM strategy. CRM and CRSM have the potential to be general plasmid-based gene mutagenesis tools.

Improved bacterial recombineering by parallelized protein discovery.

Exploiting bacteriophage-derived homologous recombination processes has enabled precise, multiplex editing of microbial genomes and the construction of billions of customized genetic variants in a single day. The techniques that enable this, multiplex automated genome engineering (MAGE) and directed evolution with random genomic mutations (DIvERGE), are however, currently limited to a handful of microorganisms for which single-stranded DNA-annealing proteins (SSAPs) that promote efficient recombineering have been identified. Thus, to enable genome-scale engineering in new hosts, efficient SSAPs must first be found. Here we introduce a high-throughput method for SSAP discovery that we call "serial enrichment for efficient recombineering" (SEER). By performing SEER in Escherichia coli to screen hundreds of putative SSAPs, we identify highly active variants PapRecT and CspRecT. CspRecT increases the efficiency of single-locus editing to as high as 50% and improves multiplex editing by 5- to 10-fold in E. coli, while PapRecT enables efficient recombineering in Pseudomonas aeruginosa, a concerning human pathogen. CspRecT and PapRecT are also active in other, clinically and biotechnologically relevant enterobacteria. We envision that the deployment of SEER in new species will pave the way toward pooled interrogation of genotype-to-phenotype relationships in previously intractable bacteria.

Experimental Confirmation that an Uncommon rrs Gene Mutation (g878a) of Mycobacterium tuberculosis Confers Resistance to Streptomycin.

The effective treatment of patients diagnosed with drug-resistant tuberculosis is highly dependent on the ability to rapidly and accurately determine the antibiotic susceptibility profile of the Mycobacterium tuberculosis isolate(s) involved. Thus, as more clinical microbiology laboratories advance toward the use of DNA sequence-based diagnostics, it is imperative that their predictive functions extend beyond the well-known resistance mutations in order to also encompass as many of the lower-frequency mutations as possible. However, in most cases, fundamental experimental proof that links these uncommon mutations with phenotypic resistance is lacking. One such example is the g878a polymorphism within the rrs 16S rRNA gene. We, and others, have identified this mutation within a small number of drug-resistant isolates, although a consensus regarding exactly which aminoglycoside antibiotic(s) it confers resistance to has not previously been reached. Here, we have employed oligonucleotide-mediated recombineering to introduce the g878a polymorphism into the rrs gene of Mycobacterium bovis BCG, a close relative of M. tuberculosis, and demonstrate that it confers low-level resistance to streptomycin alone. It does not confer cross-resistance to amikacin, capreomycin, or kanamycin. We also demonstrate that the rrsg878a mutation exerts a substantial fitness defect in vitro that may at least in part explain why clinical isolates bearing this mutation appear to be quite rare. Overall, this study provides clarity to the phenotype attributable to the rrsg878a mutation and is relevant to the future implementation of genomics-based diagnostics as well as the clinical management of patients in whom this particular polymorphism is encountered.

Deletion of FRT-sites by no-SCAR recombineering in Escherichia coli.

Lambda-Red recombineering is the most commonly used method to create point mutations, insertions or deletions in Escherichia coli and other bacteria, but usually an Flp recognition target (FRT) scar-site is retained in the genome. Alternative scarless recombineering methods, including CRISPR/Cas9-assisted methods, generally require cloning steps and/or complex PCR schemes for specific targeting of the genome. Here we describe the deletion of FRT scar-sites by the scarless Cas9-assisted recombineering method no-SCAR using an FRT-specific guide RNA, sgRNAFRT, and locus-specific ssDNA oligonucleotides. We applied this method to construct a scarless E. coli strain suitable for gradual induction by l-arabinose. Genome sequencing of the resulting strain and its parent strains demonstrated that no additional mutations were introduced along with the simultaneous deletion of two FRT scar-sites. The FRT-specific no-SCAR selection by sgRNAFRT/Cas9 may be generally applicable to cure FRT scar-sites of E. coli strains constructed by classical λ-Red recombineering.

Contributions of the Human Cytomegalovirus UL30-Associated Open Reading Frames to Infection.

Transposon-based insertional mutagenesis screens have assessed how disruption of numerous human cytomegalovirus (HCMV) open reading frames (ORFs) impacts in vitro viral replication. Insertional mutagenesis of the HCMV UL30 gene was previously found to substantially inhibit production of viral progeny. However, there are a number of putative UL30-associated ORFs, and it is unclear how they impact viral replication. Here, we report on the contributions of the eight UL30-associated ORFs to infection. We find that deletion of the canonically annotated UL30 ORF substantially reduces production of infectious virus at both high and low multiplicities of infection (MOI). This deletion likely has complex effects on viral replication, as we find that it reduces the expression of neighboring non-UL30-associated ORFs. Mutation of the initiating methionine of the canonical UL30 ORF indicated that it is dispensable for high- and low-MOI infection in the highly passaged AD169 strain, although it is important for low-MOI infection in the less-passaged TB40/E strain. Comutation of eight methionines in the UL30 region results in a low-MOI viral replication defect, as does mutation of the TATA box responsible for the most abundant UL30 transcript, which is found to be necessary for the accumulation of multiple UL30-associated protein isoforms during infection. In total, our data indicate the importance of the UL30-associated ORFs during low-MOI HCMV infection and further highlight the difficulty associated with the functional interrogation of broadly disruptive mutations: e.g., large deletions or transposon insertions.IMPORTANCE Viral genes and their products are the critical determinants of viral infection. Human cytomegalovirus (HCMV) encodes many gene products whose roles during viral infection have not been assessed. Elucidation of the contributions that various HCMV gene products make to infection provides insight into the infectious program, which could potentially be used to limit HCMV-associated morbidity, a major issue during congenital infection and in immunosuppressed populations. Here, we explored the role of HCMV's UL30-associated gene products and found that they are important for HCMV replication. Future work elucidating the mechanisms through which they contribute to viral infection could highlight novel avenues for therapeutic intervention.

Development of a New Recombineering System for Agrobacterium Species.

The discovery of new and efficient genetic engineering technologies for Agrobacterium will broaden the capacity for fundamental research on this genus and its utilization as a transgenic vehicle. In this study, we aim to develop an efficient recombineering system for Agrobacterium species. We examined isolates of Agrobacterium and the closely related genus Rhizobium to identify pairs of ET-like recombinases that would aid in the recombineering of Agrobacterium species. Four pairs of ET-like recombinases, named RecETh1h2h3h4AGROB6, RecETh1h2P3RHI597, RecETRHI145, and RecEThRHI483, were identified in Agrobacterium tumefaciens strain B6, Rhizobium leguminosarum bv. trifolii WSM597, Rhizobium sp. strain LC145, and Rhizobium sp. strain Root483D2, respectively. Eight more candidate recombineering systems were generated by combining the new ET-like recombinases with Redγ or Pluγ. The PluγETRHI145 system, the RecETh1h2h3h4AGROB6 system, and the PluγEThRHI483 system were determined to be the most efficient recombineering systems for the type strains A. tumefaciens C58, A. tumefaciens EHA105, and Rhizobium rhizogenes NBRC 13257, respectively. The utility of these systems was demonstrated by knocking out the istB-istA fusion gene in C58, the celI gene in EHA105, and the 3'-to-5' exonuclease gene and endoglucanase gene in NBRC 13257. Our work provides an effective genetic manipulation strategy for Agrobacterium species. IMPORTANCEAgrobacterium is a powerful transgenic vehicle for the genetic manipulation of numerous plant and fungal species and even animal cells. In addition to improving the utility of Agrobacterium as a transgenic vehicle, genetic engineering tools are important for revealing crucial components that are functionally involved in transfer DNA (T-DNA) translocation events. This work developed an efficient and versatile recombineering system for Agrobacterium. The successful genome modification of Agrobacterium strains revealed that this new recombineering system could be used for the genetic engineering of Agrobacterium.

Recombineering using RecET-like recombinases from Xenorhabdus and its application in mining of natural products.

Xenorhabdus can produce a large number of secondary metabolites with insecticidal, bacteriostatic, and antitumor activities. Efficient gene editing tools will undoubtedly facilitate the functional genomics research and bioprospecting in Xenorhabdus. In this study, BlastP analysis using the amino acid sequences of Redαß or RecET recombinases as queries resulted in the identification of an operon (XBJ1_operon 0213) containing RecET-like recombinases encoding genes from the genome of Xenorhabdus bovienii strain SS-2004. Three proteins encoded by this operon was indispensable for full activity of recombineering, namely XBJ1-1173 (RecE-like protein), XBJ1-1172 (RecT-like protein), and XBJ1-1171 (single-strand annealing protein). Using this newly developed recombineering system, a gene cluster responsible for biosynthesis of a novel secondary metabolite (Min16) was identified from X. stockiae HN_xs01 strain. Min16 which exhibited antibacterial and cytotoxic activities was determined to be a cyclopeptide composed of Acyl-Phe-Thr-Phe-Pro-Pro-Leu-Val by using high-resolution mass spectrometry and nuclear magnetic resonance analysis, and was designated as changshamycin. This host-specific recombineering system was proven to be effective for gene editing in Xenorhabdus, allowing for efficient discovery of novel natural products with attractive bioactivities. KEY POINTS: • Screening and identification of efficient gene editing tools from Xenorhabdus • Optimization of the Xenorhabdus electroporation parameters • Discovery of a novel cyclopeptide compound with multiple biological activities.

RNAi expression tuning, microfluidic screening, and genome recombineering for improved protein production in Saccharomyces cerevisiae.

The cellular machinery that supports protein synthesis and secretion lies at the foundation of cell factory-centered protein production. Due to the complexity of such cellular machinery, the challenge in generating a superior cell factory is to fully exploit the production potential by finding beneficial targets for optimized strains, which ideally could be used for improved secretion of other proteins. We focused on an approach in the yeast Saccharomyces cerevisiae that allows for attenuation of gene expression, using RNAi combined with high-throughput microfluidic single-cell screening for cells with improved protein secretion. Using direct experimental validation or enrichment analysis-assisted characterization of systematically introduced RNAi perturbations, we could identify targets that improve protein secretion. We found that genes with functions in cellular metabolism (YDC1, AAD4, ADE8, and SDH1), protein modification and degradation (VPS73, KTR2, CNL1, and SSA1), and cell cycle (CDC39), can all impact recombinant protein production when expressed at differentially down-regulated levels. By establishing a workflow that incorporates Cas9-mediated recombineering, we demonstrated how we could tune the expression of the identified gene targets for further improved protein production for specific proteins. Our findings offer a high throughput and semirational platform design, which will improve not only the production of a desired protein but even more importantly, shed additional light on connections between protein production and other cellular processes.

[Editing of Phage Genomes - Recombineering-Assisted SpCas9 Modification of Model Coliphages T7, T5, and T3].

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

CRISPR-derived genome editing technologies for metabolic engineering.

In metabolic engineering, genome editing tools make it much easier to discover and evaluate relevant genes and pathways and construct strains. Clustered regularly interspaced palindromic repeats (CRISPR)-associated (Cas) systems now have become the first choice for genome engineering in many organisms includingindustrially relevant ones. Targeted DNA cleavage by CRISPR-Cas provides variousgenome engineering modes such as indels, replacements, large deletions, knock-in and chromosomal rearrangements, while host-dependent differences in repair pathways need to be considered. The versatility of the CRISPR system has given rise to derivative technologies that complement nuclease-based editing, which causes cytotoxicity especially in microorganisms. Deaminase-mediated base editing installs targeted point mutations with much less toxicity. CRISPRi and CRISPRa can temporarily control gene expression without changing the genomic sequence. Multiplex, combinatorial and large scale editing are made possible by streamlined design and construction of gRNA libraries to further accelerates comprehensive discovery, evaluation and building of metabolic pathways. This review summarizes the technical basis and recent advances in CRISPR-related genome editing tools applied for metabolic engineering purposes, with representative examples of industrially relevant eukaryotic and prokaryotic organisms.

RecT Recombinase Expression Enables Efficient Gene Editing in Enterococcus spp.

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

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

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

Optimizing recombineering in Corynebacterium glutamicum.

Owing to the increasing demand for amino acids and valuable commodities that can be produced by Corynebacterium glutamicum, there is a pressing need for new rapid genome engineering tools that improve the speed and efficiency of genomic insertions, deletions, and mutations. Recombineering using the λ Red system in Escherichia coli has proven very successful at genetically modifying this organism in a quick and efficient manner, suggesting that optimizing a recombineering system for C. glutamicum will also improve the speed for genomic modifications. Here, we maximized the recombineering efficiency in C. glutamicum by testing the efficacy of seven different recombinase/exonuclease pairs for integrating single-stranded DNA and double-stranded DNA (dsDNA) into the genome. By optimizing the homologous arm length and the amount of dsDNA transformed, as well as eliminating codon bias, a dsDNA recombineering efficiency of 13,250 transformed colonies/109 viable cells was achieved, the highest efficiency currently reported in the literature. Using this optimized system, over 40,000 bp could be deleted in one transformation step. This recombineering strategy will greatly improve the speed of genetic modifications in C. glutamicum and assist other systems, such as clustered regularly interspaced short palindromic repeats and multiplexed automated genome engineering, in improving targeted genome editing.

Identification of phage recombinase function unit in genus Corynebacterium.

Phage recombinase function unit (PRFU) plays a key role in the life cycle of phage. Repurposing this system such as lambda-Redαß or Rac-RecET for recombineering has gained success in Escherichia coli. Previous studies have showed that most PRFUs only worked well in its native hosts but poorly in the distant species. Thus, identification of new PRFUs in specific species is necessary for the development of its corresponding genetic engineering tools. Here, we present a thorough study of PRFUs in the genomes of genus Corynebacterium. We first used a database to database searching method to facilitate accurate prediction of novel PRFUs in 423 genomes. A total number of 60 sets of unique PRFUs were identified and divided into 8 types based on evolution affinities. Recombineering ability of the 8 representative PRFUs was experimentally verified in the Corynebacterium glutamicum ATCC 13032 strain. In particular, PRFU from C. aurimucosum achieved highest efficiency in both ssDNA and dsDNA mediated recombineering, which is expected to greatly facilitate genome engineering in genus Corynebacterium. These results will provide new insights for the study and application of PRFUs. KEY POINTS: • First report of bioinformatic mining and systematic analysis of Phage recombinase function unit (PRFU) in Corynebacterium genomes. • Recombineering ability of the representative PRFUs was experimentally verified in Corynebacterium glutamicum ATCC 13032 strain. • PRFU with the highest recombineering efficiency at 10-2 magnitude was identified from Corynebacterium aurimucosum.

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

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

Gene modification by fast-track recombineering for cellular localization and isolation of components of plant protein complexes.

To accelerate the isolation of plant protein complexes and study cellular localization and interaction of their components, an improved recombineering protocol is described for simple and fast site-directed modification of plant genes in bacterial artificial chromosomes (BACs). Coding sequences of fluorescent and affinity tags were inserted into genes and transferred together with flanking genomic sequences of desired size by recombination into Agrobacterium plant transformation vectors using three steps of E. coli transformation with PCR-amplified DNA fragments. Application of fast-track recombineering is illustrated by the simultaneous labelling of CYCLIN-DEPENDENT KINASE D (CDKD) and CYCLIN H (CYCH) subunits of kinase module of TFIIH general transcription factor and the CDKD-activating CDKF;1 kinase with green fluorescent protein (GFP) and mCherry (green and red fluorescent protein) tags, and a PIPL (His18 -StrepII-HA) epitope. Functionality of modified CDKF;1 gene constructs is verified by complementation of corresponding T-DNA insertion mutation. Interaction of CYCH with all three known CDKD homologues is confirmed by their co-localization and co-immunoprecipitation. Affinity purification and mass spectrometry analyses of CDKD;2, CYCH, and DNA-replication-coupled HISTONE H3.1 validate their association with conserved TFIIH subunits and components of CHROMATIN ASSEMBLY FACTOR 1, respectively. The results document that simple modification of plant gene products with suitable tags by fast-track recombineering is well suited to promote a wide range of protein interaction and proteomics studies.

Donor plasmids for phenotypically neutral chromosomal gene insertions in Enterobacteriaceae.

Recombineering using bacteriophage lambda Red recombinase (λ-Red) uses homologous recombination to manipulate bacterial genomes and is commonly applied to disrupt genes to elucidate their function. This is often followed by the introduction of a wild-type copy of the gene on a plasmid to complement its function. This is often not, however, at a native copy number and the introduction of a chromosomal version of a gene can be a desirable solution to provide wild-type copy expression levels of an allele in trans. Here, we present a simple methodology based on the λ-Red-based 'gene doctoring' technique, where we developed tools used for chromosomal tagging in a conserved locus downstream of glmS and found no impact on a variety of important phenotypes. The tools described provide an easy, quick and inexpensive method of chromosomal modification for the creation of a library of insertion mutants to study gene function.