Efficient Genetic Modification Research Articles

CRISPR-based gene editing in plants: Focus on reagents and their delivery tools

Introduction: CRISPR-Cas9 technology has revolutionized plant genome editing, providing precise and efficient methods for genetic modification. This study focuses on the advancements and delivery of CRISPR-Cas9 in plant gene editing. Methods: A comprehensive search in scientific databases, including PubMed, ScienceDirect, and Google Scholar, was conducted to gather information on CRISPR-Cas9 gene editing and its delivery in precise gene modification in plants. Results: The evolving landscape of CRISPR nucleases has led to the development of innovative technologies, enhancing plant research. However, successful editing is contingent on efficient delivery of genome engineering reagents. CRISPR-based gene editing in plants utilizes diverse delivery methods: Agrobacterium-mediated transformation for bacterial transfer, biolistic transformation for physical gene insertion, electroporation for direct gene entry, expression of developmental regulators for gene expression modulation, and tobacco rattle virus as a viral vector, each offering distinct advantages for precise and efficient genetic modification in plants. Conclusion: CRISPR-Cas9 gene editing stands as a pivotal advancement in plant genetics, offering precise gene manipulation with applications in agriculture and biotechnology. The continuous refinement of reagent delivery tools reinforces CRISPR-Cas9's transformative role in plant genome editing, with significant implications for broader scientific applications.

CRISPR-Cas9 mediated genome editing of Kluyveromyces marxianus for iterative, multiplexed gene disruption and pathway integration.

Kluyveromyces marxianus, a thermotolerant, fast-growing, Crabtree-negative yeast, is a promising chassis for the manufacture of various bioproducts. Although several genome editing tools are available for this yeast, these tools still require refinement to enable more convenient and efficient genetic modification. In this study, we engineered the K. marxianus NBRC 104275 strain by impairing the nonhomologous end joining and enhancing the homologous recombination machinery, which resulted in improved homology-directed repair effective on homology arms of up to 40 bp in length. Additionally, we simplified the CRISPR-Cas9 editing system by constructing a strain for integrative expression of Cas9 nuclease and plasmids bearing different selection markers for gRNA expression, thereby facilitating iterative genome editing without the need for plasmid curing. We demonstrated that tRNA was more effective than the hammerhead ribozyme for processing gRNA primary transcripts, and readily assembled tRNA-gRNA arrays were used for multiplexed editing of at least four targets. This editing tool was further employed for simultaneous scarless in vivo assembly of a 12-kb cassette from three fragments and marker-free integration for expressing a fusion variant of fatty acid synthase, as well as the integration of genes for starch hydrolysis. Together, the genome editing tool developed in this study makes K. marxianus more amenable to genetic modification and will facilitate more extensive engineering of this nonconventional yeast for chemical production.

Streptococcus pyogenes Cas9 ribonucleoprotein delivery for efficient, rapid and marker-free gene editing in Trypanosoma and Leishmania.

Kinetoplastids are unicellular eukaryotic flagellated parasites found in a wide range of hosts within the animal and plant kingdoms. They are known to be responsible in humans for African sleeping sickness (Trypanosoma brucei), Chagas disease (Trypanosoma cruzi), and various forms of leishmaniasis (Leishmania spp.), as well as several animal diseases with important economic impact (African trypanosomes, including Trypanosoma congolense). Understanding the biology of these parasites necessarily implies the ability to manipulate their genomes. In this study, we demonstrate that transfection of a ribonucleoprotein complex, composed of recombinant Streptococcus pyogenes Cas9 (SpCas9) and an invitro-synthesized guide RNA, results in rapid and efficient genetic modifications of trypanosomatids, in marker-free conditions. This approach was successfully developed to inactivate, delete, and mutate candidate genes in various stages of the life cycle of T. brucei and T. congolense, and Leishmania promastigotes. The functionality of SpCas9 in these parasites now provides, to the research community working on these parasites, a rapid and efficient method of genome editing, without requiring plasmid construction and selection by antibiotics but requires only cloning and PCR screening of the clones. Importantly, this approach is adaptable to any wild-type parasite.

Molecular Marvels: Small Molecules Paving the Way for Enhanced Gene Therapy.

In the rapidly evolving landscape of genetic engineering, the advent of CRISPR-Cas technologies has catalyzed a paradigm shift, empowering scientists to manipulate the genetic code with unprecedented accuracy and efficiency. Despite the remarkable capabilities inherent to CRISPR-Cas systems, recent advancements have witnessed the integration of small molecules to augment their functionality, introducing new dimensions to the precision and versatility of gene editing applications. This review delves into the synergy between CRISPR-Cas technologies based specifically on Cas9 and small-molecule drugs, elucidating the pivotal role of chemicals in optimizing target specificity and editing efficiency. By examining a diverse array of applications, ranging from therapeutic interventions to agricultural advancements, we explore how the judicious use of chemicals enhances the precision of CRISPR-Cas9-mediated genetic modifications. In this review, we emphasize the significance of small-molecule drugs in fine-tuning the CRISPR-Cas9 machinery, which allows researchers to exert meticulous control over the editing process. We delve into the mechanisms through which these chemicals bolster target specificity, mitigate off-target effects, and contribute to the overall refinement of gene editing outcomes. Additionally, we discuss the potential of chemical integration in expanding the scope of CRISPR-Cas9 technologies, enabling tailored solutions for diverse genetic manipulation challenges. As CRISPR-Cas9 technologies continue to evolve, the integration of small-molecule drugs emerges as a crucial avenue for advancing the precision and applicability of gene editing techniques. This review not only synthesizes current knowledge but also highlights future prospects, paving the way for a deeper understanding of the synergistic interplay between CRISPR-Cas9 systems and chemical modulators in the pursuit of more controlled and efficient genetic modifications.

A Review on Citric Acid Production by Yarrowia lipolytica Yeast: Past and Present Challenges and Developments

The biosynthesis of citric acid (CA) and its derivatives is of great interest due to its wide range of applications in various manufacturing sectors. The fungus Aspergillus niger is mainly used for the commercial production of CA, using sucrose and molasses as the primary carbon sources. Since the 1960s, intensive research has been underway to introduce Yarrowia lipolytica yeast as an alternative to traditional fungal technology. This review discusses the practical uses of CA and its derivatives. Also, the challenges and developments that have led to efficient and green CA synthesis technologies using Y. lipolytica are outlined. The nutrient medium requirements and the use of various carbon sources, encompassing pure substrates and industry, agriculture, and food waste are considered. Additionally, the choice and improvement of strain producers, including efficient mutagenesis, genetic modification, and screening methods, are discussed.

Development of a CRISPR/Cas9D10A Nickase (nCas9)-Mediated Genome Editing Tool in Streptomyces.

Streptomycetes have a strong ability to produce a vast array of bioactive natural products (NPs) widely used in agriculture and veterinary/human medicine. The recently developed CRISPR/Cas9-based genome editing tools have greatly facilitated strain improvement for target NP overproduction as well as novel NP discovery in Streptomyces. However, CRISPR/Cas9 shows high toxicity to the host, limiting its application in many Streptomyces strains with a low DNA transformation efficiency. In this study, we developed a low-toxicity CRISPR/Cas9D10A nickase (nCas9)-based genome editing tool in the model strain Streptomyces coelicolor M145. We showed that in the presence of both targeting sgRNA and Cas proteins, utilization of nCas9 instead of Cas9 significantly reduced the toxicity to the host and greatly enhanced cell survival. Using this tool, we achieved deletion of single genes and gene clusters with efficiencies of 87-100 and 63-87%, and simultaneous deletion of two genes or gene clusters with efficiencies of 47 and 43%, respectively. The editing efficiency of nCas9 is comparable to that of the Cas9-mediated editing tool. Finally, the nCas9-based editing tool was successfully applied for genome editing in the industrial rapamycin-producing strain Streptomyces rapamycinicus, in which CRISPR/Cas9 cannot work well. We achieved the deletion of three tested genes with an efficiency of 27.2-30%. Collectively, the CRISPR/nCas9-based editing tool offers a convenient and efficient genetic modification system for the engineering of streptomycetes, particularly those with low DNA transformation efficiency.

Liposome-Based Carriers for CRISPR Genome Editing.

The CRISPR-based genome editing technology, known as clustered regularly interspaced short palindromic repeats (CRISPR), has sparked renewed interest in gene therapy. This interest is accompanied by the development of single-guide RNAs (sgRNAs), which enable the introduction of desired genetic modifications at the targeted site when used alongside the CRISPR components. However, the efficient delivery of CRISPR/Cas remains a challenge. Successful gene editing relies on the development of a delivery strategy that can effectively deliver the CRISPR cargo to the target site. To overcome this obstacle, researchers have extensively explored non-viral, viral, and physical methods for targeted delivery of CRISPR/Cas9 and a guide RNA (gRNA) into cells and tissues. Among those methods, liposomes offer a promising approach to enhance the delivery of CRISPR/Cas and gRNA. Liposomes facilitate endosomal escape and leverage various stimuli such as light, pH, ultrasound, and environmental cues to provide both spatial and temporal control of cargo release. Thus, the combination of the CRISPR-based system with liposome delivery technology enables precise and efficient genetic modifications in cells and tissues. This approach has numerous applications in basic research, biotechnology, and therapeutic interventions. For instance, it can be employed to correct genetic mutations associated with inherited diseases and other disorders or to modify immune cells to enhance their disease-fighting capabilities. In summary, liposome-based CRISPR genome editing provides a valuable tool for achieving precise and efficient genetic modifications. This review discusses future directions and opportunities to further advance this rapidly evolving field.

Therapeutic applications of CRISPR/Cas9 gene editing technology for the treatment of ocular diseases.

Ocular diseases are a highly heterogeneous group of phenotypes, caused by a spectrum of genetic variants and environmental factors that exhibit diverse clinical symptoms. As a result of its anatomical location, structure and immune privilege, the eye is an ideal system to assess and validate novel genetic therapies. Advances in genome editing have revolutionized the field of biomedical science, enabling researchers to understand the biology behind disease mechanisms and allow the treatment of several health conditions, including ocular pathologies. The advent of clustered regularly interspaced short palindromic repeats (CRISPR)-based gene editing facilitates efficient and specific genetic modifications in the nucleic acid sequence, resulting in permanent changes at the genomic level. This approach has advantages over other treatment strategies and is promising for the treatment of various genetic and non-genetic ocular conditions. This review provides an overview of the CRISPR/CRISPR-associated protein 9 (Cas9) system and summarizes recent advances in the therapeutic application of CRISPR/Cas9 for the treatment of various ocular pathologies, as well as future challenges.

Proximal binding of dCas9 at a DNA double strand break stimulates homology-directed repair as a local inhibitor of classical non-homologous end joining.

In CRISPR/Cas9 genome editing, the tight and persistent target binding of Cas9 provides an opportunity for efficient genetic and epigenetic modification on genome. In particular, technologies based on catalytically dead Cas9 (dCas9) have been developed to enable genomic regulation and live imaging in a site-specific manner. While post-cleavage target residence of CRISPR/Cas9 could alter the pathway choice in repair of Cas9-induced DNA double strand breaks (DSBs), it is possible that dCas9 residing adjacent to a break may also determine the repair pathway for this DSB, providing an opportunity to control genome editing. Here, we found that loading dCas9 onto a DSB-adjacent site stimulated homology-directed repair (HDR) of this DSB by locally blocking recruitment of classical non-homologous end-joining (c-NHEJ) factors and suppressing c-NHEJ in mammalian cells. We further repurposed dCas9 proximal binding to increase HDR-mediated CRISPR genome editing by up to 4-fold while avoiding exacerbation of off-target effects. This dCas9-based local inhibitor provided a novel strategy of c-NHEJ inhibition in CRISPR genome editing in place of small molecule c-NHEJ inhibitors, which are often used to increase HDR-mediated genome editing but undesirably exacerbate off-target effects.

Bacillus integrative plasmid system combining a synthetic gene circuit for efficient genetic modifications of undomesticated Bacillus strains

BackgroundOwing to CRISPR-Cas9 and derivative technologies, genetic studies on microorganisms have dramatically increased. However, the CRISPR-Cas9 system is still difficult to utilize in many wild-type Bacillus strains owing to Cas9 toxicity. Moreover, less toxic systems, such as cytosine base editors, generate unwanted off-target mutations that can interfere with the genetic studies of wild-type strains. Therefore, a convenient alternative system is required for genetic studies and genome engineering of wild-type Bacillus strains. Because wild-type Bacillus strains have poor transformation efficiencies, the new system should be based on broad-host-range plasmid-delivery systems.ResultsHere, we developed a Bacillus integrative plasmid system in which plasmids without the replication initiator protein gene (rep) of Bacillus are replicated in a donor Bacillus strain by Rep proteins provided in trans but not in Bacillus recipients. The plasmids were transferred to recipients through a modified integrative and conjugative element, which is a wide host range plasmid-delivery system. Genetic mutations were generated in recipients through homologous recombination between the transferred plasmid and the genome. The system was improved by adding a synthetic gene circuit for efficient screening of the desired mutations by double crossover recombination in recipient strains. The improved system exhibited a mutation efficiency of the target gene of approximately 100% in the tested wild-type Bacillus strains.ConclusionThe Bacillus integrative plasmid system developed in this study can generate target mutations with high efficiency when combined with a synthetic gene circuit in wild-type Bacillus strains. The system is free of toxicity and unwanted off-target mutations as it generates the desired mutations by traditional double crossover recombination. Therefore, our system could be a powerful tool for genetic studies and genome editing of Cas9-sensitive wild-type Bacillus strains.

In vivo killing of primary HIV-infected cells by peripheral-injected early memory-enriched anti-HIV duoCAR T cells.

HIV-specific chimeric antigen receptor-T cell (CAR T cell) therapies are candidates to functionally cure HIV infection in people with HIV (PWH) by eliminating reactivated HIV-infected cells derived from latently infected cells within the HIV reservoir. Paramount to translating such therapeutic candidates successfully into the clinic will require anti-HIV CAR T cells to localize to lymphoid tissues in the body and eliminate reactivated HIV-infected cells such as CD4+ T cells and monocytes/macrophages. Here we show that i.v. injected anti-HIV duoCAR T cells, generated using a clinical-grade anti-HIV duoCAR lentiviral vector, localized to the site of active HIV infection in the spleen of humanized mice and eliminated HIV-infected PBMCs. CyTOF analysis of preinfusion duoCAR T cells revealed an early memory phenotype composed predominantly of CCR7+ stem cell-like/central memory T cells (TSCM/TCM) with expression of some effector-like molecules. In addition, we show that anti-HIV duoCAR T cells effectively sense and kill HIV-infected CD4+ T cells and monocytes/macrophages. Furthermore, we demonstrate efficient genetic modification of T cells from PWH on suppressive ART into anti-HIV duoCAR T cells that subsequently kill autologous PBMCs superinfected with HIV. These studies support the safety and efficacy of anti-HIV duoCAR T cell therapy in our presently open phase I/IIa clinical trial (NCT04648046).

Rapid and efficient generation of antigen-specific isogenic T cells from cryopreserved blood samples.

Clustered regularly interspaced short palindromic repeats/CRISPR‐associated protein 9 (CRISPR/Cas9)‐mediated gene editing has been leveraged for the modification of human and mouse T cells. However, limited experience is available on the application of CRISPR/Cas9 electroporation in cryopreserved T cells collected during clinical trials. To address this, we aimed to optimize a CRISPR/Cas9‐mediated gene editing protocol compatible with peripheral blood mononuclear cells (PBMCs) samples routinely produced during clinical trials. PBMCs from healthy donors were used to generate knockout T‐cell models for interferon‐γ, Cbl proto‐oncogene B (CBLB), Fas cell surface death receptor (Fas) and T‐cell receptor (TCRαβ) genes. The effect of CRISPR/Cas9‐mediated gene editing on T cells was evaluated using apoptosis assays, cytokine bead arrays and ex vivo and in vitro stimulation assays. Our results demonstrate that CRISPR/Cas9‐mediated gene editing of ex vivo T cells is efficient and does not overtly affect T‐cell viability. Cytokine release and T‐cell proliferation were not affected in gene‐edited T cells. Interestingly, memory T cells were more susceptible to CRISPR/Cas9 gene editing than naïve T cells. Ex vivo and in vitro stimulation with antigens resulted in equivalent antigen‐specific T‐cell responses in gene‐edited and untouched control cells, making CRISPR/Cas9‐mediated gene editing compatible with clinical antigen‐specific T‐cell activation and expansion assays. Here, we report an optimized protocol for rapid, viable and highly efficient genetic modification in ex vivo human antigen‐specific T cells, for subsequent functional evaluation and/or expansion. Our platform extends CRISPR/Cas9‐mediated gene editing for use in gold‐standard clinically used immune‐monitoring pipelines and serves as a starting point for development of analogous approaches, such as those including transcriptional activators and/or epigenetic modifiers.

Genetic Modification of Limbs Using Ex Vivo Machine Perfusion.

Genetic engineering is a promising tool to repair genetic disorders, improve graft function, or reduce immune responses toward allografts. Ex vivo organ perfusion systems have the potential to mitigate ischemic-reperfusion injury, prolong preservation time, or even rescue organ function. We aim at combining both technologies to develop a modular platform allowing the genetic modification of vascularized composite (VC) allografts. Rat hind limbs were perfused ex vivo under subnormothermic conditions with lentiviral vectors. Specific perfusion conditions such as controlled pressure, temperature, and flow rates were optimized to support the genetic modification of the limbs. Genetic modification was detected in vascular, muscular, and dermal limb tissues. Remarkably, skin follicular and interfollicular keratinocytes, as well as endothelial cells showed stable transgene expression. Furthermore, levels of injury markers such as lactate, myoglobin, and lactate dehydrogenase, as well as histological analyses showed that ex vivo limb perfusion with lentiviral vectors did not cause tissue damage and limb cytokine secretion signatures were not significantly affected. The use of ex vivo VC perfusion in combination with lentiviral vectors allows an efficient and stable genetic modification representing a robust platform to genetically engineer limbs toward increasing graft survival after transplantation.

Efficient and Cost Effective Electroporation Method to Study Primary Cilium-Dependent Signaling Pathways in the Granule Cell Precursor

The primary cilium is a critical signaling organelle found on nearly every cell that transduces Hedgehog (Hh) signaling stimuli from the cell surface. In the granule cell precursor (GCP), the primary cilium acts as a pivotal signaling center that orchestrates precursor cell proliferation by modulating the Hh signaling pathway. The investigation of primary cilium-dependent Hh signaling machinery is facilitated by in vitro genetic manipulation of the pathway components to visualize their dynamic localization to the primary cilium. However, transfection of transgenes in the primary cultures of GCPs using the currently known electroporation methods is generally costly and often results in low cell viability and undesirable transfection efficiency. This paper introduces an efficient, cost-effective, and simple electroporation protocol that demonstrates a high transfection efficiency of ~80-90% and optimal cell viability. This is a simple, reproducible, and efficient genetic modification method that is applicable to the study of the primary cilium-dependent Hedgehog signaling pathway in primary GCP cultures.

Molecular Tools to Exploit the Biotechnological Potential of Brettanomyces bruxellensis: A Review

The Brettanomyces bruxellensis species plays various roles in both the industrial and food sectors. At the biotechnological level, B. bruxellensis is considered to be a promising species for biofuel production. Its presence in alcoholic beverages can be detrimental or beneficial to the final product; B. bruxellensis can contribute to spoilage of wine and beer, but can also produce good aromas. However, little is known about its genetic characteristics and, despite the complete sequencing of several B. bruxellensis genomes and knowledge of its metabolic pathways, the toolkits for its efficient and easy genetic modification are still underdeveloped. Moreover, the different ploidy states and the high level of genotype diversity within this species makes the development of effective genetic manipulation tools challenging. This review summarizes the available tools for the genetic manipulation of B. bruxellensis and how they may be employed to improve the quality of wine and beer.

Optimized Transgene Delivery Using Third-Generation Lentiviruses.

The lentivirus system enables efficient genetic modification of both dividing and non‐dividing cells and therefore is a useful tool for elucidating developmental processes and disease pathogenesis. The development of third‐generation lentiviruses has resulted in improved biosafety, low immunogenicity, and substantial packaging capabilities. However, because third‐generation lentiviruses require successful co‐transfection with four plasmids, this typically means that lower titers are attained. This is problematic, as it is often desirable to produce purified lentiviruses with high titers (>1 × 108 TU/ml), especially for in vivo applications. The manufacturing process for lentiviruses involves several critical experimental factors that can influence titer, purity, and transduction efficiency. Here, we describe a straightforward, stepwise protocol for the reproducible manufacture of high‐titer third‐generation lentiviruses (1 × 108 to 1 × 109 TU/ml). This optimized protocol enhances transgene expression by use of Lipofectamine transfection and optimized serum replacement medium, a single ultracentrifugation step, use of a sucrose cushion, and addition of a histone deacetylation inhibitor. Furthermore, we provide alternate methods for titration analyses, including functional and genomic integration analyses, using common laboratory techniques such as FACS as well as genomic DNA extraction and qPCR. These optimized methods will be beneficial for investigating developmental processes and disease pathogenesis in vitro and in vivo. © 2020 The Authors. Basic Protocol 1: Lentivirus production Support Protocol: Lentivirus concentration Basic Protocol 2: Lentivirus titration Alternate Protocol 1: Determination of viral titration by FACS analysis Alternate Protocol 2: Determination of viral titration by genome integration analysis

Efficient callus induction and a temperature condition for flowering and seed setting in kenaf <i>Hibiscus cannabinus</i>

Kenaf, Hibiscus cannabinus, is a fiber-enriched plant belonging to Malvaceae and is an important fiber crop. The features of kenaf of being fast-growing and fiber-enriched suggest the potential for the use of kenaf in biomass and materials. Here, we modified procedures for regeneration from kenaf explants in order to establish efficient genetic modification for improving the property of kenaf as a material. We tested several tissues of kenaf seedlings for callus induction and subsequent shoot regeneration by supplying several combinations of plant growth factors. We show the cotyledons to be the best tissue for efficient callus induction, and the rate for callus induction reached ∼98% on an improved callus inducing medium. We also show the efficient regeneration of kenaf from cotyledon-derived calli achieved by shoot induction on a shoot-inducing medium. We also achieved seed setting of the regenerated kanaf plants under a regulated growth condition in a chamber. Our efficient regeneration method and seed setting condition will enable the production of stably transformed kenaf that can improve the properties of kenaf as a material.

Vectofusin-1 Improves Transduction of Primary Human Cells with Diverse Retroviral and Lentiviral Pseudotypes, Enabling Robust, Automated Closed-System Manufacturing.

Cell and gene therapies are finally becoming viable patient treatment options, with both T cell- and hematopoietic stem cell (HSC)-based therapies being approved to market in Europe. However, these therapies, which involve the use of viral vector to modify the target cells, are expensive and there is an urgent need to reduce manufacturing costs. One major cost factor is the viral vector production itself, therefore improving the gene modification efficiency could significantly reduce the amount of vector required per patient. This study describes the use of a transduction enhancing peptide, Vectofusin-1®, to improve the transduction efficiency of primary target cells using lentiviral and gammaretroviral vectors (LV and RV) pseudotyped with a variety of envelope proteins. Using Vectofusin-1 in combination with LV pseudotyped with viral glycoproteins derived from baboon endogenous retrovirus, feline endogenous virus (RD114), and measles virus (MV), a strongly improved transduction of HSCs, B cells and T cells, even when cultivated under low stimulation conditions, could be observed. The formation of Vectofusin-1 complexes with MV-LV retargeted to CD20 did not alter the selectivity in mixed cell culture populations, emphasizing the precision of this targeting technology. Functional, ErbB2-specific chimeric antigen receptor-expressing T cells could be generated using a gibbon ape leukemia virus (GALV)-pseudotyped RV. Using a variety of viral vectors and target cells, Vectofusin-1 performed in a comparable manner to the traditionally used surface-bound recombinant fibronectin. As Vectofusin-1 is a soluble peptide, it was possible to easily transfer the T cell transduction method to an automated closed manufacturing platform, where proof of concept studies demonstrated efficient genetic modification of T cells with GALV-RV and RD114-RV and the subsequent expansion of mainly central memory T cells to a clinically relevant dose.

CReasPy-Cloning: A Method for Simultaneous Cloning and Engineering of Megabase-Sized Genomes in Yeast Using the CRISPR-Cas9 System.

Over the past decade, a new strategy was developed to bypass the difficulties to genetically engineer some microbial species by transferring (or "cloning") their genome into another organism that is amenable to efficient genetic modifications and therefore acts as a living workbench. As such, the yeast Saccharomyces cerevisiae has been used to clone and engineer genomes from viruses, bacteria, and algae. The cloning step requires the insertion of yeast genetic elements in the genome of interest, in order to drive its replication and maintenance as an artificial chromosome in the host cell. Current methods used to introduce these genetic elements are still unsatisfactory, due either to their random nature (transposon) or the requirement for unique restriction sites at specific positions (TAR cloning). Here we describe the CReasPy-cloning, a new method that combines both the ability of Cas9 to cleave DNA at a user-specified locus and the yeast's highly efficient homologous recombination to simultaneously clone and engineer a bacterial chromosome in yeast. Using the 0.816 Mbp genome of Mycoplasma pneumoniae as a proof of concept, we demonstrate that our method can be used to introduce the yeast genetic element at any location in the bacterial chromosome while simultaneously deleting various genes or group of genes. We also show that CReasPy-cloning can be used to edit up to three independent genomic loci at the same time with an efficiency high enough to warrant the screening of a small (<50) number of clones, allowing for significantly shortened genome engineering cycle times.

Targeted knock-in into the OVA locus of chicken cells using CRISPR/Cas9 system with homology-independent targeted integration

It is anticipated that transgenic avian species will be used as living bioreactors for the production of biopharmaceutical proteins. Precise tissue-specific expression of exogenous genes is a major challenge for the development of avian bioreactors. No robust vector is currently available for highly efficient and specific expression. In recent years, genome-editing techniques such as the CRISPR/Cas9 system have emerged as efficient and user-friendly genetic modification tools. Here, to apply the CRISPR/Cas9 system for the development of transgenic chickens, guide RNA sequences (gRNAs) of the CRISPR/Cas9 system for the ovalbumin (OVA) locus were evaluated for the oviduct-specific expression of exogenous genes. An EGFP gene expression cassette was introduced into the OVA locus of chicken DF-1 and embryonic fibroblasts using the CRISPR/Cas9 system mediated by homology-independent targeted integration. For the knock-in cells, EGFP expression was successfully induced by activation of the endogenous OVA promoter using the dCas9-VPR transactivation system. The combination of gRNAs designed around the OVA TATA box was important to induce endogenous OVA gene expression with high efficiency. These methods provide a useful tool for studies on the creation of transgenic chicken bioreactors and the activation of tissue-specific promoters.