DelsGate, a robust and rapid gene deletion construction method.

With the increasing availability of fungal genome sequences there is great demand for fast, simple high-throughput methods to generate constructs for gene deletion. Here we describe a method that combines PCR and Gateway cloning technology together with use of the I-SceI homing endonuclease to generate precise deletion constructs in a very simple, universal and robust manner in just 2 days. These constructs are then used to produce deletion mutants in the organism of interest following applicable methods for that species. In establishing this protocol we determined empirically that 1 kb was a suitable flank length to facilitate homologous recombination in our species of interest, Ustilago maydis. The method, which we have named DelsGate (Deletions via Gateway), consists of standard PCR of only the 5' and 3' 1 kb gene flanks directly followed by in vitro Gateway cloning and final generation of the circular deletion construct by in vivo recombination in Escherichia coli. For use in DelsGate we have modified a Gateway cloning vector to include selectable markers for transformation of Ascomycetes and the Basidiomycete fungus U. maydis which causes corn smut disease. We have tested the reproducibility of the DelsGate approach by generating deletion constructs for 12 U. maydis genes. Although not tested here, the PCR and transformation steps of DelsGate should be well suited for high-throughput approaches to gene deletion construction in fungal species. DelsGate has the potential to be universal for all organisms with efficient transformation and homologous recombination systems.     

DelsGate, a robust and rapid gene deletion construction method

With the increasing availability of fungal genome sequences there is great demand for fast, simple high-throughput methods to generate constructs for gene deletion. Here we describe a method that combines PCR and Gateway cloning technology together with use of the I-SceI homing endonuclease to generate precise deletion constructs in a very simple, universal and robust manner in just 2 days. These constructs are then used to produce deletion mutants in the organism of interest following applicable methods for that species. In establishing this protocol we determined empirically that 1 kb was a suitable flank length to facilitate homologous recombination in our species of interest, Ustilago maydis. The method, which we have named DelsGate (Deletions via Gateway), consists of standard PCR of only the 5' and 3' 1 kb gene flanks directly followed by in vitro Gateway cloning and final generation of the circular deletion construct by in vivo recombination in Escherichia coli. For use in DelsGate we have modified a Gateway cloning vector to include selectable markers for transformation of Ascomycetes and the Basidiomycete fungus U. maydis which causes corn smut disease. We have tested the reproducibility of the DelsGate approach by generating deletion constructs for 12 U. maydis genes. Although not tested here, the PCR and transformation steps of DelsGate should be well suited for high-throughput approaches to gene deletion construction in fungal species. DelsGate has the potential to be universal for all organisms with efficient transformation and homologous recombination systems.     

Genome-wide Gene Deletions in Streptococcus sanguinis by High Throughput PCR

Transposon mutagenesis and single-gene deletion are two methods applied in genome-wide gene knockout in bacteria ^1,2. Although transposon mutagenesis is less time consuming, less costly, and does not require completed genome information, there are two weaknesses in this method: (1) the possibility of a disparate mutants in the mixed mutant library that counter-selects mutants with decreased competition; and (2) the possibility of partial gene inactivation whereby genes do not entirely lose their function following the insertion of a transposon. Single-gene deletion analysis may compensate for the drawbacks associated with transposon mutagenesis. To improve the efficiency of genome-wide single gene deletion, we attempt to establish a high-throughput technique for genome-wide single gene deletion using Streptococcus sanguinis as a model organism. Each gene deletion construct in S. sanguinis genome is designed to comprise 1-kb upstream of the targeted gene, the aphA-3 gene, encoding kanamycin resistance protein, and 1-kb downstream of the targeted gene. Three sets of primers F1/R1, F2/R2, and F3/R3, respectively, are designed and synthesized in a 96-well plate format for PCR-amplifications of those three components of each deletion construct. Primers R1 and F3 contain 25-bp sequences that are complementary to regions of the aphA-3 gene at their 5'' end. A large scale PCR amplification of the aphA-3 gene is performed once for creating all single-gene deletion constructs. The promoter of aphA-3 gene is initially excluded to minimize the potential polar effect of kanamycin cassette. To create the gene deletion constructs, high-throughput PCR amplification and purification are performed in a 96-well plate format. A linear recombinant PCR amplicon for each gene deletion will be made up through four PCR reactions using high-fidelity DNA polymerase. The initial exponential growth phase of S. sanguinis cultured in Todd Hewitt broth supplemented with 2.5% inactivated horse serum is used to increase competence for the transformation of PCR-recombinant constructs. Under this condition, up to 20% of S. sanguinis cells can be transformed using ~50 ng of DNA. Based on this approach, 2,048 mutants with single-gene deletion were ultimately obtained from the 2,270 genes in S. sanguinis excluding four gene ORFs contained entirely within other ORFs in S. sanguinis SK36 and 218 potential essential genes. The technique on creating gene deletion constructs is high throughput and could be easy to use in genome-wide single gene deletions for any transformable bacteria.     

基于PCR的质粒快速定点诱变方法建立与评价

基因突变对生命的进化具有重要意义,针对质粒的DNA定点诱变技术也是基因工程、蛋白质工程研究中的重要手段之一。为了提高质粒定点诱变的效率,本研究利用引物部分重叠的设计方案,使用Tm值相对固定的引物设计模式,将同一引物分为重叠区（Tm=50±2℃）和非重叠区（Tm=60±2℃）,并在严格控制模板用量（2 pg/kb）的基础上,通过20个PCR循环对目标质粒进行定点诱变扩增,随后取0.5 μL产物直接用于转化。在FastPfu Fly酶系中,利用此法构建了6个含碱基替换、缺失和插入的质粒,均获得成功,突变效率可达96%以上,阳性克隆获得数达70个以上。此外,利用4种不同PCR酶系对该法的适用性进行了评价,结果表明突变效率均可达93%以上,阳性克隆获得数均在10个以上。通过适当增加PCR模板用量（10 pg/kb）并使用纯化后的PCR产物进行转化,该法可适用于转化效率大于106（cfu/μg）的任意感受态细胞,对应的突变效率可大于91%,阳性克隆获得数大于20。根据本法的作用原理,该方案适合质粒中10～20 bp（因重叠区GC含量及碱基序列的不同而改变）以内的任意碱基替换和插入,以及任意长度的DNA片段缺失。且具有通用性强、耗时少、诱变成功率高、成本低、对感受态及转化效率无特殊要求等优点,适合各实验室的日常研究使用。     

DNA oligonucleotide-assisted genetic manipulation increases transformation and homologous recombination efficiencies: Evidence from gene targeting of Dictyostelium discoideum

Artificial gene alteration by homologous recombination in living cells, termed gene targeting, presents fundamental and considerable knowledge of in vivo gene function. In principle, this method can possibly be applied to any type of genes and transformable cells. However, its success is limited due to a low frequency of homologous recombination between endogenous targeted gene and exogenous transgene. Here, we describe a general gene-targeting method in which co-transformation of DNA oligonucleotides (oligomers) could significantly increase the homologous recombination frequency and transformation efficiency. The oligomers were simply designed such that they were identical to both the ends of the homologous flanking regions of the targeting construct. Using this strategy, both targeted alleles of diploid cells were simultaneously replaced in a single transformation procedure. Thus, the simplicity and versatility of this method applicable to any type of cell may increase the application of gene targeting.     

Size Of Gene Specific Inverted Repeat - Dependent Gene Deletion In Saccharomyces cerevisiae

We describe here an approach for rapidly producing scar-free and precise gene deletions in S. cerevisiae with high efficiency. Preparation of the disruption gene cassette in this approach was simply performed by overlap extension-PCR of an invert repeat of a partial or complete sequence of the targeted gene with URA3. Integration of the prepared disruption gene cassette to the designated position of a target gene leads to the formation of a mutagenesis cassette within the yeast genome, which consists of a URA3 gene flanked by the targeted gene and its inverted repeat between two short identical direct repeats. The inherent instability of the inverted sequences in close proximity facilitates the self-excision of the entire mutagenesis cassette deposited in the genome and promotes homologous recombination resulting in a seamless deletion via a single transformation. This rapid assembly circumvents the difficulty during preparation of disruption gene cassettes composed of two inverted repeats of the URA3, which requires the engineering of unique restriction sites for subsequent digestion and T4 DNA ligation in vitro. We further identified that the excision of the entire mutagenesis cassette flanked by two DRs in the transformed S. cerevisiae is dependent on the length of the inverted repeat of which a minimum of 800 bp is required for effective gene deletion. The deletion efficiency improves with the increase of the inverted repeat till 1.2 kb. Finally, the use of gene-specific inverted repeats of target genes enables simultaneous gene deletions. The procedure has the potential for application on other yeast strains to achieve precise and efficient removal of gene sequences.     

One step construction of Agrobacterium-Recombination-ready-plasmids (OSCAR), an efficient and robust tool for ATMT based gene deletion construction in fungi

Increasing availability of genomic data and sophistication of analytical methodology in fungi has elevated the need for functional genomics tools in these organisms. Previously we reported a method called DelsGate for rapid preparation of deletion constructs for protoplast-mediated fungal transformation systems, which is based on Gateway? technology. However, over the past several years Agrobacteriumtumefaciens-mediated transformation (ATMT) has become the preferred genetic transformation method for an increasing number of fungi. Therefore, we developed a method for One Step Construction of Agrobacterium-Recombination-ready-plasmids (OSCAR), to rapidly create deletion constructs for ATMT systems. The OSCAR methodology involves PCR amplification of the upstream and downstream flanks of the gene of interest, using gene specific primers each with a 5' extension containing one of four different attB recombination sites, modified from the Invitrogen MultiSite Gateway? system. Amplified gene flanks are then mixed with specifically designed marker and binary vectors and treated with BP clonase, generating the deletion construct in a single cloning step. The entire process of deletion construct preparation can be accomplished in just 2days. Using OSCAR we generated eight targeted deletion constructs and used two of them to generate deletion mutants in Verticillium dahliae by ATMT. In summary, OSCAR methodology combines PCR and Gateway? technology to rapidly and robustly generate precise deletion constructs for fungal ATMT and homologous gene replacement.     

A QuikChange-like method to realize efficient blunt-ended DNA directional cloning and site-directed mutagenesis simultaneously

Here we present a QuikChange-like method to efficiently realize blunt-ended DNA cloning and conveniently introduce a site-directed mutation to recombinant plasmid at the same time. After blunt-ended DNA ligation and transformation, the plasmid DNA mixture is extracted from pooled transformants and directly used as template for PCR amplification with a pair of complementary mutagenic primers. With this method, sam1 gene was inserted into pUC19 vector by blunt-end ligation, and a unique restriction site Spe I was introduced to the recombinant plasmid at the same time. The randomly selected transformants were analyzed by DNA sequencing, and most of the clones were found to have correct sequences. However, no correct construct was found from randomly selected transformants after traditional blunt-ended DNA ligation and transformation.     

Insertion-Duplication Mutagenesis in Streptococcus pneumoniae: Targeting Fragment Length Is a Critical Parameter in Use as a Random Insertion Tool

To examine whether insertion-duplication mutagenesis with chimeric DNA as a transformation donor could be valuable as a gene knockout tool for genomic analysis in Streptococcus pneumoniae, we studied the transformation efficiency and targeting specificity of the process by using a nonreplicative vector with homologous targeting inserts of various sizes. Insertional recombination was very specific in targeting homologous sites. While the recombination rate did not depend on which site or region was targeted, it did depend strongly on the size of the targeting insert in the donor plasmid, in proportion to the fifth power of its length for inserts of 100 to 500 bp. The dependence of insertion-duplication events on the length of the targeting homology was quite different from that for linear allele replacement and places certain limits on the design of mutagenesis experiments. The number of independent pneumococcal targeting fragments of uniform size required to knock out any desired fraction of the genes in a model genome with a defined probability was calculated from these data by using a combinatorial theory with simplifying assumptions. The results show that efficient and thorough mutagenesis of a large part of the pneumococcal genome should be practical when using insertion-duplication mutagenesis.     

Efficient generation of gene knockout plasmids for Dictyostelium discoideum using one-step cloning

The amoeba Dictyostelium discoideum is a well-established model organism for studying numerous aspects of cellular and developmental functions. Its rather small (~34Mb) chromosomal genome and the high efficiency of gene disruption by homologous recombination have enabled researchers to dissect various specific gene functions. We describe here the use of one-step cloning for the fast and efficient generation of deletion vectors that are produced in a one-step reaction by inserting two PCR products into an organism-specific, generic acceptor system. This worked efficiently for all 16 tested constructs directed against genes in the amoeba Dictyostelium discoideum. Saving cost and time, the used protocol represents a significant advancement in the generation of such plasmids compared to the conventionally applied restriction enzyme/ligation approach. Using appropriate selection markers, similar systems could also be useful in other organisms, where genes can be knocked out by homologous recombination.     

A rapid and simple method for inactivating chromosomal genes in Yersinia

A polymerase chain reaction (PCR)-based procedure without any cloning step was developed for a rapid mutagenesis/deletion of chromosomal target genes in Yersinia. For this purpose, a PCR fragment carrying an antibiotic resistance gene flanked by regions homologous to the target locus is electroporated into a recipient strain expressing the highly proficient homologous recombination system encoded by plasmid pKOBEG-sacB. Two PCR procedures were tested to generate an amplification product formed of an antibiotic resistance gene flanked by short (55 bp) or long (500 bp) homology extensions. Using this method, three chromosomal loci were successfully disrupted in Yersinia pseudotuberculosis. The use of this technique allows rapid and efficient large-scale mutagenesis of Yersinia target chromosomal genes.     

Gene conversion of immunoglobulin variable regions in mutagenesis cassettes by replacement PCR mutagenesis.

A technique, Replacement PCR Mutagenesis, was developed to replace one immunoglobulin variable region (V) in a M13 phage cassette with a different, homologous V. This allows the use of the same mutagenesis and subsequent expression vectors for many V regions or V segments. The method combines PCR of V fragments and in vitro mutagenesis. Primers homologous to 3' and 5' ends of both V regions initiate PCR synthesis of the V DNA fragment (donor) that will replace the V region (recipient) in M13. Donor V PCR DNA may originate from mRNA, cloned V genes or genomic templates. The donor V PCR DNA is denatured and annealed to the M13 cassette containing the recipient V to be supplanted. The second strand is synthesized, transfected into bacteria and mutant plaques selected by hybridization. Since restriction sites in primers are not required, altered primer-encoded amino acids are avoided. Further, the PCR donor piece can be of any length if it shares homology with the recipient gene. This allows construction and expression of complete gene replacements and chimeras. This method is also applicable to V "humanization" and studying sets of homologous genes containing polymorphic or evolutionary disparities. The potential uses of the technique are discussed.     

Site-directed mutagenesis by combination of homologous recombination and DpnI digestion of the plasmid template in Escherichia coli

A rapid site-directed mutagenesis strategy using homologous recombination and DpnI digestion of the template in Escherichia coli is described. Briefly, inverse polymerase chain reaction amplification of the entire circular plasmid was performed by mutagenic primers with overlapping sequences ( approximately 15 bp) for generating PCR products with approximately 15 bp of homology on the terminal ends. On direct transformation of the amplified PCR products into restriction endonuclease DpnI-expressing E. coli BUNDpnI, homologous recombination occurs in E. coli while the original templates are removed via DpnI digestion in vivo, thus yielding clones harboring mutated circular plasmids. Nearly 100% efficiency was attained when this strategy was used to modify DNA sequences.     

A PCR-based system for highly efficient generation of gene replacement mutants in<Emphasis Type="Italic"> Ustilago maydis</Emphasis>

Ustilago maydis, the causative agent of corn smut disease, is one of the most versatile model systems for the study of plant pathogenic fungi. With the availability of the complete genomic sequence there is an increasing need to improve techniques for the generation of deletion mutants in order to elucidate the functions of unknown genes. Here a method is presented which allows one to generate constructs for gene replacement without the need for cloning. The 5 and 3-regions of the target gene are first amplified by PCR, and subsequently ligated directionally to a marker cassette via two distinct Sfi I sites, providing the flanking homologies needed for homologous recombination in U. maydis. Then the ligation product is used as a template for the amplification of the deletion construct, which can be used directly for transformation of U. maydis. The use of the fragments generated by PCR drastically increases the frequency of homologous recombination when compared to the linearized plasmids routinely used for gene replacement in U. maydis.Communicated by G. Jürgens     

A site-directed mutagenesis method particularly useful for creating otherwise difficult-to-make mutants and alanine scanning

Site-directed mutagenesis has become routine in molecular biology. However, many mutants can still be very difficult to create. Complicated chimerical mutations, tandem repeats, inverted sequences, GC-rich regions, and/or heavy secondary structures can cause inefficient or incorrect binding of the mutagenic primer to the target sequence and affect the subsequent amplification. In theory, these problems can be avoided by introducing the mutations into the target sequence using mutagenic fragments and so removing the need for primer-template annealing. The cassette mutagenesis uses the mutagenic fragment in its protocol; however, in most cases it needs to perform two rounds of mutagenic primer-based mutagenesis to introduce suitable restriction enzyme sites into templates and is not suitable for routine mutagenesis. Here we describe a highly efficient method in which the template except the region to be mutated is amplified by polymerase chain reaction (PCR) and the type IIs restriction enzyme-digested PCR product is directly ligated with the mutagenic fragment. Our method requires no assistance of mutagenic primers. We have used this method to create various types of difficult-to-make mutants with mutagenic frequencies of nearly 100%. Our protocol has many advantages over the prevalent QuikChange method and is a valuable tool for studies on gene structure and function.     

利用Red重组系统快速构建基因打靶载体

基因敲除小鼠模型是在哺乳动物体内研究基因功能最可靠的方法之一。利用常规的分子克隆的方法构建基因打靶载体往往工作周期长,对于难度特别大的基因有时甚至无法完成打靶载体的构建。通过合理应用Red重组系统和低拷贝中间载体,利用50bp的同源重组序列直接从BAC载体中克隆了长片段的小鼠基因组序列;将得到的基因组序列再次通过重组和改造,构建了Gpr56等基因的完全敲除并带有报告基因的打靶载体,实现了打靶载体的快速构建。     

Extending homologous sequence based on the single gene mutants by one-step PCR for efficient multiple gene knockouts

Multiple gene knockouts play an important role in metabolic engineering. The flanked homology length, homologous to the region adjacent to the target gene, of the knockout fragments has a great effect on the efficiency of multiple gene knockouts, whereas the existing gene knockout methods can only supply a very short homology. This article presents a strategy of easily extending homologous sequence based on the available strain library through one-step PCR amplification (the one-step PCR method). In this approach, the library of single gene mutants was used as the templates for PCR to amplify knockout fragments. Thus, the flanked homology can be extended as long as possible by designing primers upstream and downstream far from the target gene. Based on the one-step PCR method, we studied the effect of the homology length and the number of mutations on the efficiency of multiple gene knockouts. Our results indicated that the one-step PCR method permitted rapid and efficient construction of multiple mutants continuously or simultaneously, and a length of 200–300 bp homologous sequence was equal for multiple gene knockouts.     

Identification of a locus involved in meningococcal lipopolysaccharide biosynthesis by deletion mutagenesis

A novel method for insertion/deletion mutagenesis in meningococci was devised. This consisted of ligating a digest of total chromosomal DNA to a 1.1 kb restriction fragment containing an erythromycin-resistance marker ( ermC ), and subsequent transformation of the ligation mixture into the homologous meningococcal strain H44/76. Southern blotting of a number of the resulting erythromycin-resistant transformants demonstrated that all carried the ermC gene inserted at different positions in the chromosome. Mutants with a specific phenotype were identified by screening with the anti-lipopolysaccharide (LPS) monoclonal antibody MN4A8B2, which is specific for immunotype L3. In this way, two independent L3-negative mutant strains were isolated. In transformation experiments with chromosomal DNA from these mutants, erythromycin-resistance and lack of MN4A8B2 reactivity were always linked, showing that the insertion/deletion was in a locus involved in LPS biosynthesis. On SDS–PAGE, the mutant LPS displayed an electrophoretic mobility intermediate between that produced by the previously isolated galE and rfaF mutant strains. Chemical analysis of the mutant LPS revealed that the structure was probably lipid A–(KDO)₂–(Hep)₂. Chromosomal DNA flanking the ermC insertion in these two mutant strains was cloned, and used as probe for the isolation of the corresponding region of the wild-type strain. From hybridization and polymerase chain reaction (PCR) analysis, it could be concluded that both mutations map to the same locus. The affected gene probably encodes the glycosyltransferase necessary for adding N -acetylglucosamine to heptose.     

Mutagenesis by imprecise excision of the piggyBac transposon in Drosophila melanogaster

Mutagenesis by transposon-mediated imprecise excision is the most extensively used technique for mutagenesis in Drosophila. Although P-element is the most widely used transposon in Drosophila to generate deletion mutants, it is limited by the insertion coldspots in the genome where P-elements are rarely found. The piggyBac transposon was developed as an alternative mutagenic vector for mutagenesis of non-P-element targeted genes in Drosophila because the piggyBac transposon can more randomly integrate into the genome. Previous studies suggested that the piggyBac transposon always excises precisely from the insertion site without initiating a deletion or leaving behind an additional footprint. This unique characteristic of the piggyBac transposon facilitates reversible gene-transfer in several studies, such as the generation of induced pluripotent stem (iPS) cells from fibroblasts. However, it also raised a potential limitation of its utility in generating deletion mutants in Drosophila. In this study, we report multiple imprecise excisions of the piggyBac transposon at the sepiapterin reductase (SR) locus in Drosophila. Through imprecise excision of the piggyBac transposon inserted in the 5'-UTR of the SR gene, we generated a hypomorphic mutant allele of the SR gene which showed markedly decreased levels of SR expression. Our finding suggests that it is possible to generate deletion mutants by piggyBac transposon-mediated imprecise excision in Drosophila. However, it also suggests a limitation of piggyBac transposon-mediated reversible gene transfer for the generation of induced pluripotent stem (iPS) cells.