Homology Arms Research Articles

Two-Step Bacterial Artificial Chromosome (BAC) Engineering: Cloning of the A and B Homology Arms into the Shuttle Vector.

This protocol describes the preparation of the shuttle vector before its introduction into bacterial artificial chromosome (BAC) host cells for BAC two-step engineering. The homology arm sequences, prepared previously, are introduced by ligation into the digested shuttle vector DNA to provide sites for recombination within the BAC clone. Crude lysates of individual bacterial transformants serve as templates in polymerase chain reaction (PCR) analysis to confirm the presence of the homology arms in the recombinant shuttle vector.

Two-Step Bacterial Artificial Chromosome (BAC) Engineering: Preparation and Verification of the Recombinant Shuttle Vector.

Plasmid DNA is prepared from the recombinant shuttle vector pLD53.SCAB/A-B created by cloning of the A and B homology arms for two-step bacterial artificial chromosome (BAC) engineering. To confirm that the A-box and B-box arms have been successfully incorporated into pLD53.SCAB, the pattern of enzyme digestion of the modified plasmid is compared with that of the unmodified pLD53.SCAB. Once the shuttle vector is shown to carry the proper sequences, it is ready for transfer into the BAC host.

SPARC: Development of Pigs with mCherry‐CRE Recombinase Fusion Proteins in Cholinergic Neurons

Large animals are increasingly utilized in biomedical research and offer an advantage in that their anatomy and physiology closely parallel humans. Yet despite these advantages, rodent models dominate many areas of research, including neuroscience. This might be due in part to the greater number of technologies available to rodent researchers. Here we describe the development of pigs with mCherry and CRE‐recombinase proteins expressed in cholinergic neurons. We first cloned a 5,972 bp DNA fragment containing the predicted choline acetyltransferase (ChAT) promoter from porcine genomic DNA and placed into a pGlow TOPO™ TA expression system. We then tested for promoter activity by transfecting the expression system into porcine hippocampal slices and assessed for green fluorescent protein (GFP). We observed GFP at 60–72 hours post transfection. Confirmation of specificity was achieved by using antibody detection of ChAT. To optimize the expression system, we relied upon proven strategies used in rodents, and added an additional 535 bp of predicted promoter to the 5’ end. We then used this sequence to drive production of a mCherry‐P2A‐CRE recombinase fusion protein (ChAT‐mCherry‐P2A‐CRE). We transfected brain slice cultures from the pig hippocampus, striatum and brainstem with ChAT‐mCherry‐P2A‐CRE. Approximately 60 hours later, we isolated RNA and froze brain slices from transfected and non‐transfected samples. qRT‐PCR revealed mCherry and CRE recombinase mRNA in all transfected tissues. Expression of mCherry protein was also visible in discreet subsets of neurons in transfected samples. Studies are ongoing to confirm mCherry expression in cholinergic neurons. Key next steps include engineering our ChAT‐mCherry‐P2A‐CRE to contain GGTA1 homology arms for placement into the GGTA1 safe harbor in the pig genome, providing in vivo validation of ChAT‐mCherry‐P2A‐CRE using lentivirus pseudotyped with rabies glycoprotein, and targeting porcine fetal fibroblasts for somatic cell nuclear transfer. If successful, this model will allow for simultaneous visualization of neurons (and their innervation), and the ability to control or monitor neural activity through CRE‐mediated recombination.Support or Funding InformationHL119560, 1OT2OD026582

Investigation of the molecular biology underlying the pronounced high gene targeting frequency at the Myh9 gene locus in mouse embryonic stem cells.

The generation of genetically modified mouse models derived from gene targeting (GT) in mouse embryonic stem (ES) cells (mESCs) has greatly advanced both basic and clinical research. Our previous finding that gene targeting at the Myh9 exon2 site in mESCs has a pronounced high homologous recombination (HR) efficiency (>90%) has facilitated the generation of a series of nonmuscle myosin II (NM II) related mouse models. Furthermore, the Myh9 gene locus has been well demonstrated to be a new safe harbor for site-specific insertion of other exogenous genes. In the current study, we intend to investigate the molecular biology underlying for this high HR efficiency from other aspects. Our results confirmed some previously characterized properties and revealed some unreported observations: 1) The comparison and analysis of the targeting events occurring at the Myh9 and several widely used loci for targeting transgenesis, including ColA1, HPRT, ROSA26, and the sequences utilized for generating these targeting constructs, indicated that a total length about 6 kb with approximate 50% GC-content of the 5' and 3' homologous arms, may facilitate a better performance in terms of GT efficiency. 2) Despite increasing the length of the homologous arms, shifting the targeting site from the Myh9 exon2, to intron2, or exon3 led to a gradually reduced GT frequency (91.7, 71.8 and 50.0%, respectively). This finding provides the first evidence that the HR frequency may also be associated with the targeting site even in the same locus. Meanwhile, the decreased trend of the GT efficiency at these targeting sites was consistent with the reduced percentage of simple sequence repeat (SSR) and short interspersed nuclear elements (SINEs) in the sequences for generating the targeting constructs, suggesting the potential effects of these DNA elements on GT efficiency; 3) Our series of targeting experiments and analyses with truncated 5' and 3' arms at the Myh9 exon2 site demonstrated that GT efficiency positively correlates with the total length of the homologous arms (R = 0.7256, p<0.01), confirmed that a 2:1 ratio of the length, a 50% GC-content and the higher amount of SINEs for the 5' and 3' arms may benefit for appreciable GT frequency. Though more investigations are required, the Myh9 gene locus appears to be an ideal location for identifying HR-related cis and trans factors, which in turn provide mechanistic insights and also facilitate the practical application of gene editing.

RS-1 enhances CRISPR-mediated targeted knock-in in bovine embryos.

Targeted knock‐in (KI) can be achieved in embryos by clustered regularly interspaced short palindromic repeats (CRISPR)‐assisted homology directed repair (HDR). However, HDR efficiency is constrained by the competition of nonhomologous end joining. The objective of this study was to explore whether CRISPR‐assisted targeted KI rates can be improved in bovine embryos by exposure to the HDR enhancer RS‐1. In vitro produced zygotes were injected with CRISPR components (300 ng/µl Cas9 messenger RNA and 100 ng/µl single guide RNA against a noncoding region) and a single‐stranded DNA (ssDNA) repair template (100 ng/µl). ssDNA template contained a 6 bp XbaI site insert, allowing targeted KI detection by restriction analysis, flanked by 50 bp homology arms. Following microinjection, zygotes were exposed to 0, 3.75, or 7.5 µM RS‐1 for 24 hr. No differences were noted between groups in terms of development or genome edition rates. However, targeted KI rates were doubled in the group exposed to 7.5 µM RS‐1 compared to the others (52.8% vs. 25% and 23.1%, for 7.5, 0, and 3.75 µM, respectively). In conclusion, transient exposure to 7.5 µM RS‐1 enhances targeted KI rates resulting in approximately half of the embryos containing the intended mutation, hence allowing direct KI generation in embryos.

Homing endonuclease I-SceI-mediated Corynebacterium glutamicum ATCC 13032 genome engineering.

Corynebacterium glutamicum is widely used to produce amino acids and is a chassis for the production of value-added compounds. Effective genome engineering methods are crucial to metabolic engineering and synthetic biology studies of C. glutamicum. Herein, a homing endonuclease I-SceI-mediated genome engineering strategy was established for the model strain C. glutamicum ATCC 13032. A vegetative R6K replicon-based, suicide plasmid was employed. The plasmid, pLS3661, contains both tightly regulated, IPTG (isopropyl-β-D-1-thiogalactopyranoside)-inducible I-SceI expression elements and two I-SceI recognition sites. Following cloning of the homologous arms into pLS3661 and transfer the recombinant vector into C. glutamicum ATCC 13032, through the homologous recombination between the cloned fragment and its chromosomal allele, a merodiploid was selected under kanamycin selection. Subsequently, a merodiploid was resolved by double-stranded break repair stimulated by IPTG-stimulated I-SceI expression, generating desired mutants. The protocol obviates a pre-generated strain, transfer of a second I-SceI expression plasmid, and there is not any strain, medium, and temperature restrictions. We validated the approach via deletions of five genes (up to ~ 13.0kb) and knock-in of one DNA fragment. Furthermore, through kanamycin resistance repair, the ssDNA recombineering parameters were optimized. We hope the highly efficient method will be helpful for the studies of C. glutamicum, and potentially, to other bacteria. KEY POINTS: • Counterselection marker I-SceI-mediated C. glutamicum genome engineering • A suicide vector contains I-SceI expression elements and its recognition sites • Gene deletions and knock-in were conducted; efficiency was as high as 90% • Through antibiotic resistance repair, ssDNA recombineering parameters were optimized.

A double-locus scarless genome editing system in Escherichia coli.

To develop a convenient double-locus scarless genome editing system in Escherichia coli, based on the type II Streptococcus pyogenes CRISPR/Cas9 and λ Red recombination cassette. A two-plasmid genome editing system was constructed. The large-sized plasmid harbors the cas9 and λ Red recombination genes (gam, bet, and exo), while the small-molecular plasmid can simultaneously express two different gRNAs (targeting genome RNAs). The recombination efficiency was tested by targeting the galK, lacZ, and dbpA genes in E. coli with ssDNA or dsDNA. Resulting concurrent double-locus recombination efficiencies were 88 ± 5.5% (point mutation), 39.7 ± 4.3% (deletion/insertion), and 57.8 ± 3.4%-58.5 ± 4.1% (mixed point and deletion/insertion mutation), depending on 30 (ssDNA) or 40bp (dsDNA) homologous side arms employed. In addition, the curing efficiency of the guide plasmid expressing gRNAs for negative selection was higher (96 ± 3% in 4h) than the help plasmid carrying cas9 and λ Red (92 ± 2% in 9h). The new editing system is convenient and efficient for simultaneous double-locus recombination in the genome and should be favorable for high-throughput multiplex genome editing in synthetic biology and metabolic engineering.

Intra-Molecular Homologous Recombination of Scarless Plasmid.

Although many methods have been reported, plasmid construction compromises transformant efficiency (number of transformants per ng of DNAs) with plasmid accuracy (rate of scarless plasmids). An efficient method is two-step PCR serving DNA amplification. An accurate method is ExnaseII cloning serving homology recombination (HR). We combine DNA amplification and HR to develop an intra-molecular HR by amplifying plasmid DNAs to contain homology 5′- and 3′-terminus and recombining the plasmid DNAs in vitro. An example was to construct plasmid pET20b-AdD. The generality was checked by constructing plasmid pET21a-AdD and pET22b-AdD in parallel. The DNAs having 30-bp homology arms were optimal for intra-molecular HR, and transformation of which created 14.2 transformants/ng and 90% scarless plasmids, more than the two-step PCR and the ExnaseII cloning. Transformant efficiency correlated with the component of nicked circular plasmid DNAs of HR products, indicating nick modification in vivo leads to scar plasmids.

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.

Targeted editing of transcriptional activator MXR1 on the Pichia pastoris genome using CRISPR/Cas9 technology.

A highly efficient and targeted clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 gene editing system was constructed for Pichia pastoris (syn Komagataella phaffii). Plasmids containing single guide RNA and the methanol expression regulator 1 (MXR1) homology arms were used to precisely edit the transcriptional activator Mxr1 on the P. pastoris genome. At the S215 amino acid position of Mxr1, one, two, and three nucleotides were precisely deleted or inserted, and S215 was also mutated to S215A via a single-base substitution. Sequencing of polymerase chain reaction (PCR) amplicons in the region spanning MXR1 showed that CRISPR/Cas9 technology enabled efficient and precise gene editing of P. pastoris. The expression levels of several of the Mxr1-targeted genes, AOX1, AOX2, DAS1, and DAS2, in strains containing the various mutated variants of MXR1, were then detected through reverse transcription PCR following induction in methanol-containing culture medium. The frameshift mutations of Mxr1 led to almost zero transcription of AOX1, DAS1, and DAS2, while that of AOX2 was reduced to 60%. For the Mxr1 S215A mutant, the transcription of AOX1, AOX2, DAS1, and DAS2 was also reduced by nearly 60%. Based on these results, it is apparent that the transcription of AOX1, DAS1, and DAS2 is exclusively regulated by Mxr1 and serine phosphorylation at Mxr1 residue 215 is not critical for this function. In contrast, the transcription of AOX2 is mainly dependent on the phosphorylation of this residue. CRISPR/Cas9 technology was, therefore, successfully applied to the targeted editing of MXR1 on the P. pastoris genome, and it provided an effective method for the study of this transcription factor and its targets.

DNA repair protein RAD51 enhances the CRISPR/Cas9-mediated knock-in efficiency in brain neurons

Gene knock-in using the CRISPR/Cas9 system can be achieved in a specific population of neurons in the mouse brain, by using in utero electroporation to introduce DNA fragments into neural progenitor cells. Using this strategy, we previously knocked-in the EGFP coding sequence into the N-terminal region of the β-actin gene specifically in the pyramidal neurons in layer 2/3 of the somatosensory cortex. However, the knock-in efficiency was less than 2% of the transfected neurons. In this study, we sought to improve the knock-in efficiency using this system. First, we varied the length of the homology arms of the β-actin donor template DNA, and found that the knock-in efficiency was increased to ∼14% by extending the length of the 5′ and 3’ homology arms to 1.6 kb and 2.0 kb, respectively. We then tested the effect of the DNA repair protein RAD51 and the knock-in efficiency was increased up to 2.5-fold when co-transfecting with two different β-actin and a camk2a targeting EGFP knock-in modules. The RAD51 overexpression did not alter the migration of developing neurons, density or morphology of the dendritic spines compared to those in neurons not transfected with RAD51. RAD51 expression will be useful for increasing the knock-in efficiency in neurons in vivo by CRISPR/Cas9-mediated homology directed repair (HDR).

CRISPR/Cas9-Mediated Integration of Large Transgene into Pig CEP112 Locus.

Clustered regularly interspaced short palindromic repeats (CRISPR)-associated protein 9 (Cas9) is a precise genome manipulating tool that can produce targeted gene mutations in various cells and organisms. Although CRISPR/Cas9 can efficiently generate gene knockout, the gene knock-in (KI) efficiency mediated by homology-directed repair remains low, especially for large fragment integration. In this study, we established an efficient method for the CRISPR/Cas9-mediated integration of large transgene cassette, which carries salivary gland-expressed multiple digestion enzymes (≈ 20 kbp) in CEP112 locus in pig fetal fibroblasts (PFFs). Our results showed that using an optimal homology donor with a short and a long arm yielded the best CRISPR/Cas9-mediated KI efficiency in CEP112 locus, and the targeting efficiency in CEP112 locus was higher than in ROSA26 locus. The CEP112 KI cell lines were used as nuclear donors for somatic cell nuclear transfer to create genetically modified pigs. We found that KI pig (705) successfully expressed three microbial enzymes (β-glucanase, xylanase, and phytase) in salivary gland. This finding suggested that the CEP112 locus supports exogenous gene expression by a tissue-specific promoter. In summary, we successfully targeted CEP112 locus in pigs by using our optimal homology arm system and established a modified pig model for foreign digestion enzyme expression in the saliva.

Modulation of Cas9 level for efficient CRISPR-Cas9-mediated chromosomal and plasmid gene deletion in Bacillus thuringiensis.

To set up an efficient gene editing system in Bacillus thuringiensis (Bt) using CRISPR-Cas9 by demonstrating deletion of chromosomal and plasmid genes. CRISPR-Cas9 from Streptococcus pyogenes was found to function in Bt cells, resulting in DNA cleavage that is lethal to the cells. The system was assessed for its ability to mediate gene editing by knock-out of the protease genes nprA (neutral protease A) and aprA (alkaline protease A). Gene editing was not detected when the Bacillus-derived pBCX was used to carry CRISPR-Cas9 elements and a DNA repair template. When the Cas9 promoter was replaced with the sporulation-specific promoter cyt2A, a Bt ∆nprA clone was obtained, but this plasmid construct did not give reproducible results. Bt ∆nprA ∆aprA and Bt ∆aprA deletion mutants were finally generated when the Lactobacillus plantarum-derived plasmid pLPPR9 was used, likely due to its lower copy number reducing Cas9 toxicity. Only three to four clones each needed to be screened to identify the desired gene-modified mutants. Conversely, efficient editing of the plasmid vip3A gene required the use of pBCX and longer homology sequences for the repair template. Capitalizing on the differential impact of plasmid copy number and homology arm length, we devised distinct yet simple and efficient approaches to chromosomal and plasmid gene deletion for Bt that condense the screening process, minimize screening, and facilitate multiple consecutive gene editing steps.

Indel locations are determined by template polarity in highly efficient in vivo CRISPR/Cas9-mediated HDR in Atlantic salmon

Precise gene editing such as CRISPR/Cas9-mediated homology directed repair (HDR) can increase our understanding of gene function and improve traits of importance for aquaculture. This fine-tuned technology has not been developed for farmed fish including Atlantic salmon. We performed knock-in (KI) of a FLAG element in the slc45a2 gene in salmon using sense (S), anti-sense (AS) and double-stranded (ds) oligodeoxynucleotide (ODN) templates with short (24/48/84 bp) homology arms. We show in vivo ODN integration in almost all the gene edited animals, and demonstrate perfect HDR rates up to 27% in individual F0 embryos, much higher than reported previously in any fish. HDR efficiency was dependent on template concentration, but not homology arm length. Analysis of imperfect HDR variants suggest that repair occurs by synthesis-dependent strand annealing (SDSA), as we show for the first time in any species that indel location is dependent on template polarity. Correct ODN polarity can be used to avoid 5′-indels interrupting the reading frame of an inserted sequence and be of importance for HDR template design in general.

Publisher Correction: An efficient gene knock-in strategy using 5'-modified double-stranded DNA donors with short homology arms.

An amendment to this paper has been published and can be accessed via a link at the top of the paper.

Genome Editing As an Approach to the Study of in Vivo Transcription Reprogramming.

CDK8-mediated transcriptional reprogramming is essential for an extensive gene expression. Constitutive knockouts of the cdk8 gene are lethal at the morula stage. For modeling transcriptional reprogramming in an adult organism, we investigated the possibility to attenuate the CDK8 kinase activity with a F97G mutation in exon 3 of the cdk8 gene. According to preliminary experimental data, this mutation should lead to a decrease in CDK8 kinase activity. To edit the genome of laboratory mice, the CRISPR/Cas9 technology was used, in which the introduction of a double-stranded gap occurred at a distance of 128 nucleotide pairs from the planned site of the introduced mutation. To introduce the mutation, a matrix for homologous repair was used as part of plasmid DNA, with homologous arms 903 and 484 bp in the 5'-3' region from the point of double-stranded rupture, respectively. As a result, mice with site-specific target mutations in exon 3 of the cdk8 gene were obtained. We for the first time demonstrated a high efficacy of the mutation 128 bp apart from the site of double-strand break. Viable animals with the F97G mutation in the catalytic domain of CDK8 kinase were obtained for the first time. The resulting cdk8 mutant mice will be used in subsequent studies to simulate the processes involving transcription reprogramming.

Genetically modified rabbits as bio-producers and biomodels

Genetically modified (GM) animals are necessary to solve the global problems of humanity related to nutrition and health. Rabbits, as laboratory, domestic and farm animals, occupy a special niche in research. GM rabbits are promising as bioreactors for producing biologically active (BA) proteins with milk or blood, and are in demand in Biomedicine as biomodels of diseases. To date, many GM rabbits-biomodels, producers of recombinant proteins have been created in the world using CRISPR/Cas9 technology. All-Russian Research Institute of Animal Physiology, Biochemistry and Nutrition has experience in obtaining transgenic rabbitsproducers of human BA proteins with milk by microinjecting recombinant DNA into zygote pronuclei. The possibility of site-specific modification of the rabbit whey acidic protein (WAP) gene using CRISPR/Cas9 technology is discussed. A DNA matrix containing homology arms to the WAP rabbit gene and site-specific CRISPR/Cas9 components in plasmid form were obtained. Microinjections of rabbit zygotes were performed and embryo survival was evaluated in vitro. The efficiency of using the green fluorescent protein gene under the cytomegalovirus promoter in the DNA matrix as an indicator of homologically directed repair was evaluated. This work can be useful for obtaining rabbits that produce with milk BA protein instead of WAP.

An efficient gene knock-in strategy using 5'-modified double-stranded DNA donors with short homology arms.

Here, we report a rapid CRISPR-Cas9-mediated gene knock-in strategy that uses Cas9 ribonucleoprotein and 5'-modified double-stranded DNA donors with 50-base-pair homology arms and achieved unprecedented 65/40% knock-in rates for 0.7/2.5 kilobase inserts, respectively, in human embryonic kidney 293T cells. The identified 5'-end modification led to up to a fivefold increase in gene knock-in rates at various genomic loci in human cancer and stem cells.

The Pediatric Acute Leukemia Fusion Oncogene ETO2‐GLIS2 Increases Self‐Renewal and Alters Differentiation in a Human Induced Pluripotent Stem Cells‐Derived Model

Supplemental Digital Content is available in the text.

Homologous recombination repair intermediates promote efficient de novo telomere addition at DNA double-strand breaks.

The healing of broken chromosomes by de novo telomere addition, while a normal developmental process in some organisms, has the potential to cause extensive loss of heterozygosity, genetic disease, or cell death. However, it is unclear how de novo telomere addition (dnTA) is regulated at DNA double-strand breaks (DSBs). Here, using a non-essential minichromosome in fission yeast, we identify roles for the HR factors Rqh1 helicase, in concert with Rad55, in suppressing dnTA at or near a DSB. We find the frequency of dnTA in rqh1Δ rad55Δ cells is reduced following loss of Exo1, Swi5 or Rad51. Strikingly, in the absence of the distal homologous chromosome arm dnTA is further increased, with nearly half of the breaks being healed in rqh1Δ rad55Δ or rqh1Δ exo1Δ cells. These findings provide new insights into the genetic context of highly efficient dnTA within HR intermediates, and how such events are normally suppressed to maintain genome stability.