High-efficiency Genome Editing Research Articles

CRISPR-based genome editing of a diurnal rodent, Nile grass rat (Arvicanthis niloticus)

BackgroundDiurnal and nocturnal mammals have evolved distinct pathways to optimize survival for their chronotype-specific lifestyles. Conventional rodent models, being nocturnal, may not sufficiently recapitulate the biology of diurnal humans in health and disease. Although diurnal rodents are potentially advantageous for translational research, until recently, they have not been genetically tractable. The present study aims to address this major limitation by developing experimental procedures necessary for genome editing in a well-established diurnal rodent model, the Nile grass rat (Arvicanthis niloticus).ResultsA superovulation protocol was established, which yielded nearly 30 eggs per female grass rat. Fertilized eggs were cultured in a modified rat 1-cell embryo culture medium (mR1ECM), in which grass rat embryos developed from the 1-cell stage into blastocysts. A CRISPR-based approach was then used for gene editing in vivo and in vitro, targeting Retinoic acid-induced 1 (Rai1), the causal gene for Smith-Magenis Syndrome, a neurodevelopmental disorder. The CRISPR reagents were delivered in vivo by electroporation using an improved Genome-editing via Oviductal Nucleic Acids Delivery (i-GONAD) method. The in vivo approach produced several edited founder grass rats with Rai1 null mutations, which showed stable transmission of the targeted allele to the next generation. CRISPR reagents were also microinjected into 2-cell embryos in vitro. Large deletion of the Rai1 gene was confirmed in 70% of the embryos injected, demonstrating high-efficiency genome editing in vitro.ConclusionWe have established a set of methods that enabled the first successful CRISPR-based genome editing in Nile grass rats. The methods developed will guide future genome editing of this and other diurnal rodent species, which will promote greater utility of these models in basic and translational research.

Low-Toxicity and High-Efficiency Streptomyces Genome Editing Tool Based on the Miniature Type V-F CRISPR/Cas Nuclease AsCas12f1.

Genome editing tools based on SpCas9 and FnCpf1 have facilitated strain improvements for natural product production and novel drug discovery in Streptomyces. However, due to high toxicity, their editing requires high DNA transformation efficiency, which is unavailable in most streptomycetes. The transformation efficiency of an all-in-one editing tool based on miniature Cas nuclease AsCas12f1 was significantly higher than those of SpCas9 and FnCpf1 in tested streptomycetes, which is due to its small size and weak DNA cleavage activity. Using this tool, in Streptomyces coelicolor, we achieved 100% efficiency for single gene or gene cluster deletion and 46.7 and 40% efficiency for simultaneous deletion of two genes and two gene clusters, respectively. AsCas12f1 was successfully extended to Streptomyces hygroscopicus SIPI-054 for efficient genome editing, in which SpCas9/FnCpf1 does not work well. Collectively, this work offers a low-toxicity, high-efficiency genome editing tool for streptomycetes, particularly those with low DNA transformation efficiency.

Non-Viral RNA-Lipid Nanoparticles for High Efficiency Genome Editing of CD34+ Hematopoietic Stem and Progenitor Cells for Advanced Cell and Gene Therapies

Background and Aims: The application of CRISPR-Cas9 gene editing in hematopoietic stem (HSC) and progenitor cells (HSPCs) has the potential to revolutionize the treatment of genetic blood disorders by correcting disease-causing mutations. Previously, we demonstrated the utility of a novel lipid nanoparticle (LNP) reagent for the multiplexed engineering of gene-edited CAR T cells, showing high cell viabilities, rapid onset editing, and potent CAR-mediated killing. In this current work, we demonstrate the capability of LNPs for the efficient and gentle delivery of genetic material to CD34+ HSPCs. The benefits of a non-viral LNP-mediated approach paves the way for the development of next generation gene therapy products in the HSC field. Methods: LNPs encapsulating S.p. Cas9 mRNA and CD33 or CD45 targeted guide RNA (sgRNA) were produced using the scalable NanoAssemblr® NxGen™ microfluidic platform technology ( Fig. 1A). The LNP composition was optimized for the gentle and efficient cargo delivery to HSC/HSPCs. Purified human CD34+ cells were cultured, stimulated, and treated by the direct addition of the RNA-LNPs. Various commercial cell sources were tested including mobilized peripheral blood and cord blood. Gene knockout and viability were assessed using flow cytometry, cell proliferation using an automated cell counter, and differentiation capacity by colony-forming unit (CFU) assays. Results: One-step addition of LNPs to CD34+ HSPCs resulted in 84 ± 6% CD33 and 81 ± 2% CD45 knockout efficiencies (n=6 CD34+ donors, Fig. 1B). After treatment, cells maintained on average 95 ± 3% absolute cell viability, compared to 99% viability of the untreated cells, Fig. 1C. Furthermore, LNP treated HSCs maintained excellent cell proliferation, with over >90% relative cell proliferation to the untreated controls. The CFU assays showed no significant change in relative lineage formation for both the RNA-LNPs and the empty LNP vehicle control ( Fig. 1D). Finally, LNP production was successfully scaled-up using microfluidics from discovery to pre-clinical scales. Conclusions: LNP-mediated CRISPR-Cas9 mRNA delivery is a promising approach for gene editing in HSPCs. The simple and gentle nature of LNP cell treatment allows for multiple genetic engineering steps for simultaneous expression and deletion of proteins for novel gene therapies. Furthermore, LNPs can be easily manufactured using microfluidics, enabling small-scale screening of RNA libraries and rapid scale-up for clinical translation.

Establishment of high-efficiency genome editing in white birch (Betula platyphylla Suk.).

Establishment of high-efficiency genome editing in white birch (Betula platyphylla Suk.).

A geminivirus crosses the monocot-dicot boundary and acts as a viral vector for gene silencing and genome editing

IntroductionMembers of the family Geminiviridae have been reported to infect either a monocot plant or a dicot plant, but not both. This study reports a geminivirus, Wheat Dwarf India Virus (WDIV), first identified in wheat, that is capable of infecting both monocot and dicot plants and acting as a viral vector. ObjectivesThis study was aimed at developing a broad host range viral vector system for reverse genetics and genome editing. MethodsHere we used a wheat isolate of WDIV and Ageratum yellow leaf curl betasatellite (AYLCB) for infectivity assays and vector development. We performed Agrobacterium-mediated inoculation of WDIV and AYLCB in wheat, oat, barley, corn, soybean, and tobacco. To examine the potential of WDIV to act as a viral vector, we modified the WDIV genome and cloned DNA fragments of the phytoene desaturase (PDS) genes from wheat and tobacco, separately. For gene editing experiments, tobacco lines expressing Cas9 were infiltrated with a WDIV-based vector carrying gRNA targeting the PDS gene. ResultsAbout 80 to 90% of plants inoculated with infectious clones of WDIV alone or WDIV together with AYLCB showed mild symptoms, whereas some plants showed more prominent symptoms. WDIV and AYLCB were detected in the systemically infected leaves of all the plant species. Furthermore, the inoculation of the WDIV vector carrying PDS fragments induced silencing of the PDS gene in both wheat and tobacco plants. We also observed high-efficiency genome editing in the Cas9-expressing tobacco plants that were inoculated with WDIV vector-carrying gRNA. ConclusionDetection of WDIV in naturally infected wheat, barley, and sugarcane in the field and its ability to systemically infect wheat, oat, barley, corn, soybean, and tobacco under laboratory conditions, provides compelling evidence that WDIV is the first geminivirus identified with the capability of infecting both monocot and dicot plant species. The wide host range of WDIV can be exploited for developing a single vector system for high-throughput genome editing in many plant species.

Developing a UV-visible reporter-assisted CRISPR/Cas9 gene editing system to alter flowering time in Chrysanthemum indicum.

Developing a UV-visible reporter-assisted CRISPR/Cas9 gene editing system to alter flowering time in Chrysanthemum indicum.

Genome Editing in Chlamydomonas reinhardtii Using Cas9-gRNA Ribonucleoprotein Complex: A Step-by-Step Guide.

Genome editing technologies have provided opportunities to manipulate literally any genomic location, opening new avenues for reverse genetics-based improvements. Among them, CRISPR/Cas9 is the most versatile tool for genome editing applications in prokaryotes and eukaryotes. Here, we provide a guide to successfully carry out high-efficiency genome editing in Chlamydomonas reinhardtii using preassembled CRISPR/Cas9-gRNA ribonucleoprotein (RNP) complexes.

High-efficiency genome editing in plants mediated by a Cas9 gene containing multiple introns

The recent discovery of the mode of action of the CRISPR/Cas9 system has provided biologists with a useful tool for generating site-specific mutations in genes of interest. In plants, site-targeted mutations are usually obtained by the stable transformation of a Cas9 expression construct into the plant genome. The efficiency of introducing mutations in genes of interest can vary considerably depending on the specific features of the constructs, including the source and nature of the promoters and terminators used for the expression of the Cas9 gene and the guide RNA, and the sequence of the Cas9 nuclease itself. To optimize the efficiency of the Cas9 nuclease in generating mutations in target genes in Arabidopsis thaliana, we investigated several features of its nucleotide and/or amino acid sequence, including the codon usage, the number of nuclear localization signals (NLSs), and the presence or absence of introns. We found that the Cas9 gene codon usage had some effect on its activity and that two NLSs worked better than one. However, the highest efficiency of the constructs was achieved by the addition of 13 introns into the Cas9 coding sequence, which dramatically improved the editing efficiency of the constructs. None of the primary transformants obtained with a Cas9 gene lacking introns displayed a knockout mutant phenotype, whereas between 70% and 100% of the primary transformants generated with the intronized Cas9 gene displayed mutant phenotypes. The intronized Cas9 gene was also found to be effective in other plants such as Nicotiana benthamiana and Catharanthus roseus.

Preclinical Evaluation for Engraftment of Gene-Edited CD34+ Cells with a Sickle Cell Disease Mutation in a Rhesus Transplantation Model

Sickle cell disease (SCD) is caused by a 20A>T mutation in the β-globin gene. State-of-the-art genome editing technologies have the potential to correct the SCD mutation in hematopoietic stem cells (HSCs), producing adult hemoglobin (Hb) while simultaneously eliminating sickle Hb. We have demonstrated efficient gene correction in SCD CD34+ cells with SCD mutation-specific guide RNA, Cas9 mRNA/protein, and single strand donor DNA, resulting in ~30% gene correction and ~50% indels at the DNA level, and ~60% normal β-globin production at the protein level in in vitro erythroid differentiation (ASH 2018). Gene correction by homology directed repair is thought to be enhanced by cell proliferation; however, cell proliferation might reduce stemness of HSCs. To investigate this hypothesis, we sought to evaluate engraftment of gene-edited CD34+ HSCs in a non-human primate model. To model SCD gene correction, a β-to-βs globin conversion was designed in rhesus macaques. Mobilized rhesus CD34+ cells (n=2) were electroporated using the GMP-compliant, FDA Master File-supported, and scalable MaxCyte GT System to deliver rhesus β-globin-targeting guide RNA (the same target site as the SCD mutation-specific guide RNA), SpCas9 protein, and single strand donor DNA including a SCD mutation (20A>T). We also added an adjuvant to improve gene conversion efficiencies. Following erythroid differentiation, gene correction efficiency was evaluated at DNA levels by deep sequencing and at protein levels by reverse-phase HPLC. We observed high-efficiency genome editing without the adjuvant (20-30% gene conversion and 61-64% indels), and further enhanced genome editing with the adjuvant (51-59% gene conversion and 36-39% indels). After erythroid differentiation, we observed production of βs-globin protein (~100%) but not normal β-globin in gene-edited cells. We then evaluated engraftment of gene-edited rhesus CD34+ cells with β-to-βs globin conversion (n=2, 13U005 and 12U011). Mobilized rhesus CD34+ cells (3.4-3.8e7) were pre-stimulated for 2 days, and edited cells were cryopreserved after electroporation with editing tools. Small aliquots of edited cells (before and after cryopreservation) were differentiated into erythroid cells in vitro, resulting in 17-26% of gene conversion and 57-71% of indels at the DNA level and 50-100% of β-globin production at the protein level, with no difference observed between aliquots taken before and after cryopreservation. Following 9.5 Gy total body irradiation, the frozen edited CD34+ cells (1.6-2.2e7) were injected into autologous macaques. We observed robust recovery of blood counts in 13U005, while peripheral blood recovery was delayed in 12U011, who was supported by serial whole blood transfusion. We observed 7-11% of gene conversion and 44-54% of indels in both granulates and lymphocytes in 13U005 1 month post-transplant. Around 15% sickle Hb production in red blood cells was detected by Hb electrophoresis in 13U005 three months post-transplant and ~7% in 12U011 two months post-transplant. Interestingly, ~10% of fetal Hb production was observed in 12U001, likely due to stress hematopoiesis. In summary, we developed a rhesus β-to-βs globin conversion model with HSC-targeted genome editing strategies. The gene-edited rhesus CD34+ cells are engraftable for at least 3 months post-transplant. Although further follow-up is necessary for transplanted animals, these findings are helpful in designing HSC-targeted gene correction trials. Figure Disclosures Li: MaxCyte, Inc: Employment. Allen:MaxCyte, Inc: Employment. Peshwa:MaxCyte, Inc: Employment.

Homologous Recombination-Based Genome Editing by Clade F AAVs Is Inefficient in the Absence of a Targeted DNA Break.

Adeno-associated virus (AAV) vectors are frequently used as donor templates for genome editing by homologous recombination. Although modification rates are typically under 1%, they are greatly enhanced by targeted double-stranded DNA breaks (DSBs). A recent report described clade F AAVs mediating high-efficiency homologous recombination-based editing in the absence of DSBs. The clade F vectors included AAV9 and a series isolated from human hematopoietic stem and progenitor cells (HSPCs). We evaluated these vectors by packaging homology donors into AAV9 and an AAVHSC capsid and examining their ability to insert GFP at the CCR5 and AAVS1 loci in human HSPCs and cell lines. As a control, we used AAV6, which effectively edits HSPCs but only when combined with a targeted DSB. Each AAV vector promoted GFP insertion in the presence of matched CCR5 or AAVS1 zinc-finger nucleases (ZFNs), but none supported detectable editing in the absence of the nucleases. Rates of editing with ZFNs correlated with transduction efficiencies for each vector, implying no differences in the ability of donor sequences delivered by the different vectors to direct genome editing. Our results, therefore, do not support that clade F AAVs can perform high-efficiency genome editing in the absence of a DSB.

Development of a Clinically Applicable Method for High-Efficiency Gene Correction of Plerixafor-Mobilized CD34+ Cells from Patients with Sickle Cell Disease

Sickle cell disease (SCD) is caused by a 20A>T mutation in the β-globin gene, and can be cured by therapeutic β-globin gene addition into hematopoietic stem cells (HSCs) with lentiviral transduction. However, this method relies upon random integration, leaving the SCD mutation intact and potentially inducing insertional mutagenesis. Genome editing technologies have the potential to correct the SCD mutation without integration, producing adult hemoglobin (Hb) while simultaneously eliminating sickle Hb. In this study, we investigated CRISPR/Cas9-based gene correction for SCD CD34+ cells.Plerixafor-mobilized SCD CD34+ cells were transfected by electroporation using the GMP-compliant, FDA Master File-supported, and scalable MaxCyte GT System to deliver SCD mutation-specific guide RNA at 200mg/ml, SpCas9 mRNA at 200mg/ml or protein at 120mg/ml, and single strand donor DNA with a normal β-globin sequence at 80, 120, or 200mg/ml. We chose Cas9 mRNA and single strand donor DNA due to the ease of clinical grade large-scale production and to avoid the need for viral vector manufacturing. Following erythroid differentiation, gene correction efficiency was evaluated at DNA levels by deep sequencing and at protein levels by reverse-phase HPLC. Cell viability was reduced to 76-87% after electroporation, compared to 90% in the control. We observed high-efficiency genome editing (29-34% gene correction and 49-58% indels) with Cas9 mRNA, showing donor DNA concentration dependence, and editing levels were comparable to Cas9 protein (39% correction and 43% indels). 15-23% Biallelic and 17-26% monoallelic gene correction were detected at the clonal level by colony assay. After erythroid differentiation, up to 54% normal β-globin production was observed with Cas9 mRNA (Figure), comparable to Cas9 protein (67%), while βs-globin amounts were markedly reduced under both conditions (6-10%). Similar correction efficiencies were obtained from two additional SCD patients' CD34+ cells at DNA levels (28-35%) and protein levels (33-56%). These data demonstrate that Cas9 mRNA and single strand donor DNA allow for efficient gene correction in SCD CD34+ cells, exceeding the therapeutic threshold of 20% in SCD.We then evaluated off-target effects on the δ-globin gene, which was reported as a major off-target site in β-globin gene editing due to high homology; however, almost no off-target effects (0.6-1.3% indels) were detected. Interestingly, gene conversion in the 9T>C polymorphism (11bp upstream of SCD mutation) on the β-globin gene was observed, and this conversion always occurred with SCD gene correction (26-33% of SCD gene correction), suggesting that gene conversion is strongly affected by distance from the target site.In addition, we evaluated genome editing among subpopulations of CD34+ cells from 3 healthy donors under the same conditions (normal β-globin to SCD mutation). We observed similar editing efficiencies (conversion and indels) among more immature (CD34+CD133+CD90+) and relatively differentiated populations (CD34+CD133+CD90-, CD34+CD133-, and CD34-) as well as among cells at different phases of the cell cycle (G0/G1, S, and G2/M), suggesting that similar gene correction efficiencies are obtained in all CD34+ cell populations, including the HSC population. We have begun efforts to evaluate gene-corrected SCD CD34+ cell engraftment in the mouse xenograft model, as similarly corrected X-CGD CD34+ cells were engrafted in immunodeficient mice.To examine the effects of indels in the β-globin gene, we next evaluated Hb production from genome-edited SCD CD34+ cells (2 patients) without donor DNA. Editing without donor DNA resulted in 63-70% indels (compared to 26-29% correction and 46-53% indels with donor DNA) and increased non-adult Hb production (small amounts of fetal Hb and significant amounts of a Hb variant), which will require further investigation to characterize.In summary, we observed efficient gene correction in SCD CD34+ cells with a simple Cas9 mRNA, single strand donor DNA, and guide RNA method, resulting in ~30% gene correction and ~50% indels. After erythroid differentiation, the majority of Hb detected was adult Hb; we detected up to 54% normal β-globin production with a marked reduction of βs-globin to ~10%. Evaluation of engraftment potential is required for gene-corrected CD34+ cells, but these methods would be clinically applicable for gene correction in SCD. [Display omitted] DisclosuresLi:MaxCyte, Inc.: Employment. Allen:MaxCyte, Inc.: Employment. Peshwa:MaxCyte, Inc.: Employment.

A genome editing vector that enables easy selection and identification of knockout cells

The CRISPR/Cas9 system is a powerful genome editing tool for disrupting the expression of specific genes in a variety of cells. However, the genome editing procedure using currently available vectors is laborious, and there is room for improvement to obtain knockout cells more efficiently. Therefore, we constructed a novel vector for high efficiency genome editing, named pGedit, which contains EGFP-Bsr as a selection marker, expression units of Cas9, and sgRNA without a terminator sequence of the U6 promoter. EGFP-Bsr is a fusion protein of EGFP and blasticidin S deaminase, and enables rapid selection and monitoring of transformants, as well as confirmation that the vector has not been integrated into the genome. By using pGedit, we targeted human ACTB, ACTG1 and mouse Nes genes coding for β-actin, γ-actin and nestin, respectively. Knockout cell lines of each gene were easily and efficiently obtained in all three cases. In this report, we show that our novel vector, pGedit, significantly facilitates genome editing.

Efficient genome editing of wild strawberry genes, vector development and validation.

SummaryThe clustered regularly interspaced short palindromic repeats (CRISPR)–Cas9 system is an effective genome editing tool for plant and animal genomes. However, there are still few reports on the successful application of CRISPR–Cas9 to horticultural plants, especially with regard to germ‐line transmission of targeted mutations. Here, we report high‐efficiency genome editing in the wild strawberry Fragaria vesca and its successful application to mutate the auxin biosynthesis gene TAA1 and auxin response factor 8 (ARF8). In our CRISPR system, the Arabidopsis U6 promoter AtU6‐26 and the wild strawberry U6 promoter FveU6‐2 were each used to drive the expression of sgRNA, and both promoters were shown to lead to high‐efficiency genome editing in strawberry. To test germ‐line transmission of the edited mutations and new mutations induced in the next generation, the progeny of the primary (T0) transgenic plants carrying the CRISPR construct was analysed. New mutations were detected in the progeny plants at a high efficiency, including large deletions between the two PAM sites. Further, T1 plants harbouring arf8 homozygous knockout mutations grew considerably faster than wild‐type plants. The results indicate that our CRISPR vectors can be used to edit the wild strawberry genome at a high efficiency and that both sgRNA design and appropriate U6 promoters contribute to the success of genomic editing. Our results open up exciting opportunities for engineering strawberry and related horticultural crops to improve traits of economic importance.

Genome editing and transcriptional repression in Pseudomonas putida KT2440 via the type II CRISPR system

BackgroundThe soil bacterium Pseudomonas putida KT2440 is a “generally recognized as safe”-certified strain with robust property and versatile metabolism. Thus, it is an ideal candidate for synthetic biology, biodegradation, and other biotechnology applications. The known genome editing approaches of Pseudomonas are suboptimal; thus, it is necessary to develop a high efficiency genome editing tool.ResultsIn this study, we established a fast and convenient CRISPR–Cas9 method in P. putida KT2440. Gene deletion, gene insertion and gene replacement could be achieved within 5 days, and the mutation efficiency reached > 70%. Single nucleotide replacement could be realized, overcoming the limitations of protospacer adjacent motif sequences. We also applied nuclease-deficient Cas9 binding at three locations upstream of enhanced green fluorescent protein (eGFP) for transcriptional inhibition, and the expression intensity of eGFP reduced to 28.5, 29.4, and 72.1% of the control level, respectively. Furthermore, based on this CRISPR–Cas9 system, we also constructed a CRISPR–Cpf1 system, which we validated for genome editing in P. putida KT2440.ConclusionsIn this research, we established CRISPR based genome editing and regulation control systems in P. putida KT2440. These fast and efficient approaches will greatly facilitate the application of P. putida KT2440.

In Vivo Base Editing of PCSK9 (Proprotein Convertase Subtilisin/Kexin Type 9) as a Therapeutic Alternative to Genome Editing.

High-efficiency genome editing to disrupt therapeutic target genes, such as PCSK9 (proprotein convertase subtilisin/kexin type 9), has been demonstrated in preclinical animal models, but there are safety concerns because of the unpredictable nature of cellular repair of double-strand breaks, as well as off-target mutagenesis. Moreover, precise knock-in of specific nucleotide changes-whether to introduce or to correct gene mutations-has proven to be inefficient in nonproliferating cells in vivo. Base editors comprising CRISPR-Cas9 (clustered regularly interspaced short palindromic repeats [CRISPR]-CRISPR-associated 9) fused to a cytosine deaminase domain can effect the alteration of cytosine bases to thymine bases in genomic DNA in a sequence-specific fashion, without the need for double-strand DNA breaks. The efficacy of base editing has not been established in vivo. The goal of this study was to assess whether in vivo base editing could be used to modify the mouse Pcsk9 gene in a sequence-specific fashion in the liver in adult mice. We screened base editors for activity in cultured cells, including human-induced pluripotent stem cells. We then delivered a base editor into the livers of adult mice to assess whether it could introduce site-specific nonsense mutations into the Pcsk9 gene. In adult mice, this resulted in substantially reduced plasma PCSK9 protein levels (>50%), as well as reduced plasma cholesterol levels (≈30%). There was no evidence of off-target mutagenesis, either cytosine-to-thymine edits or indels. These results demonstrate the ability to precisely introduce therapeutically relevant nucleotide variants into the genome in somatic tissues in adult mammals, as well as highlighting a potentially safer alternative to therapeutic genome editing.

High-Efficiency Genome Editing of Streptomyces Species by an Engineered CRISPR/Cas System.

Next-generation sequencing technologies have rapidly expanded the genomic information of numerous organisms and revealed a rich reservoir of natural product gene clusters from microbial genomes, especially from Streptomyces, the largest genus of known actinobacteria at present. However, genetic engineering of these bacteria is often time consuming and labor intensive, if even possible. In this chapter, we describe the design and construction of pCRISPomyces, an engineered Type II CRISPR/Cas system, for targeted multiplex gene deletions in Streptomyces lividans, Streptomyces albus, and Streptomyces viridochromogenes with editing efficiency ranging from 70% to 100%. We demonstrate pCRISPomyces as a powerful tool for genome editing in Streptomyces.

Rapid and highly efficient mammalian cell engineering via Cas9 protein transfection

CRISPR-Cas9 systems provide a platform for high efficiency genome editing that are enabling innovative applications of mammalian cell engineering. However, the delivery of Cas9 and synthesis of guide RNA (gRNA) remain as steps that can limit overall efficiency and ease of use. Here we describe methods for rapid synthesis of gRNA and for delivery of Cas9 protein/gRNA ribonucleoprotein complexes (Cas9 RNPs) into a variety of mammalian cells through liposome-mediated transfection or electroporation. Using these methods, we report nuclease-mediated indel rates of up to 94% in Jurkat T cells and 87% in induced pluripotent stem cells (iPSC) for a single target. When we used this approach for multigene targeting in Jurkat cells we found that two-locus and three-locus indels were achieved in approximately 93% and 65% of the resulting isolated cell lines, respectively. Further, we found that the off-target cleavage rate is reduced using Cas9 protein when compared to plasmid DNA transfection. Taken together, we present a streamlined cell engineering workflow that enables gRNA design to analysis of edited cells in as little as four days and results in highly efficient genome modulation in hard-to-transfect cells. The reagent preparation and delivery to cells is amenable to high throughput, multiplexed genome-wide cell engineering.

High-Efficiency Genome Editing and Allele Replacement in Prototrophic and Wild Strains of Saccharomyces

Current genome editing techniques available for Saccharomyces yeast species rely on auxotrophic markers, limiting their use in wild and industrial strains and species. Taking advantage of the ancient loss of thymidine kinase in the fungal kingdom, we have developed the herpes simplex virus thymidine kinase gene as a selectable and counterselectable marker that forms the core of novel genome engineering tools called the H: aploid E: ngineering and R: eplacement P: rotocol (HERP) cassettes. Here we show that these cassettes allow a researcher to rapidly generate heterogeneous populations of cells with thousands of independent chromosomal allele replacements using mixed PCR products. We further show that the high efficiency of this approach enables the simultaneous replacement of both alleles in diploid cells. Using these new techniques, many of the most powerful yeast genetic manipulation strategies are now available in wild, industrial, and other prototrophic strains from across the diverse Saccharomyces genus.

High-efficiency genome editing via 2A-coupled co-expression of fluorescent proteins and zinc finger nucleases or CRISPR/Cas9 nickase pairs.

Targeted endonucleases including zinc finger nucleases (ZFNs) and clustered regularly interspaced short palindromic repeats (CRISPRs)/Cas9 are increasingly being used for genome editing in higher species. We therefore devised a broadly applicable and versatile method for increasing editing efficiencies by these tools. Briefly, 2A peptide-coupled co-expression of fluorescent protein and nuclease was combined with fluorescence-activated cell sorting (FACS) to allow for efficient isolation of cell populations with increasingly higher nuclease expression levels, which translated into increasingly higher genome editing rates. For ZFNs, this approach, combined with delivery of donors as single-stranded oligodeoxynucleotides and nucleases as messenger ribonucleic acid, enabled high knockin efficiencies in demanding applications, including biallelic codon conversion frequencies reaching 30–70% at high transfection efficiencies and ∼2% at low transfection efficiencies, simultaneous homozygous knockin mutation of two genes with ∼1.5% efficiency as well as generation of cell pools with almost complete codon conversion via three consecutive targeting and FACS events. Observed off-target effects were minimal, and when occurring, our data suggest that they may be counteracted by selecting intermediate nuclease levels where off-target mutagenesis is low, but on-target mutagenesis remains relatively high. The method was also applicable to the CRISPR/Cas9 system, including CRISPR/Cas9 mutant nickase pairs, which exhibit low off-target mutagenesis compared to wild-type Cas9.