Transient inhibition of 53BP1 increases the frequency of targeted integration in human hematopoietic stem and progenitor cells.

Genome editing by homology directed repair (HDR) is leveraged to precisely modify the genome of therapeutically relevant hematopoietic stem and progenitor cells (HSPCs). Here, we present a new approach to increasing the frequency of HDR in human HSPCs by the delivery of an inhibitor of 53BP1 (named "i53") as a recombinant peptide. We show that the use of i53 peptide effectively increases the frequency of HDR-mediated genome editing at a variety of therapeutically relevant loci in HSPCs as well as other primary human cell types. We show that incorporating the use of i53 recombinant protein allows high frequencies of HDR while lowering the amounts of AAV6 needed by 8-fold. HDR edited HSPCs were capable of long-term and bi-lineage hematopoietic reconstitution in NSG mice, suggesting that i53 recombinant protein might be safely integrated into the standard CRISPR/AAV6-mediated genome editing protocol to gain greater numbers of edited cells for transplantation of clinically meaningful cell populations.

Genetically corrected RAG2-SCID human hematopoietic stem cells restore V(D)J-recombinase and rescue lymphoid deficiency.

ABSTRACT: Recombination-activating genes (RAG1 and RAG2) are critical for lymphoid cell development and function by initiating the variable (V), diversity (D), and joining (J) (V(D)J)-recombination process to generate polyclonal lymphocytes with broad antigen specificity. The clinical manifestations of defective RAG1/2 genes range from immune dysregulation to severe combined immunodeficiencies (SCIDs), causing life-threatening infections and death early in life without hematopoietic cell transplantation (HCT). Despite improvements, haploidentical HCT without myeloablative conditioning carries a high risk of graft failure and incomplete immune reconstitution. The RAG complex is only expressed during the G0-G1 phase of the cell cycle in the early stages of T- and B-cell development, underscoring that a direct gene correction might capture the precise temporal expression of the endogenous gene. Here, we report a feasibility study using the CRISPR/Cas9-based "universal gene-correction" approach for the RAG2 locus in human hematopoietic stem/progenitor cells (HSPCs) from healthy donors and RAG2-SCID patient. V(D)J-recombinase activity was restored after gene correction of RAG2-SCID-derived HSPCs, resulting in the development of T-cell receptor (TCR) αß and γδ CD3+ cells and single-positive CD4+ and CD8+ lymphocytes. TCR repertoire analysis indicated a normal distribution of CDR3 length and preserved usage of the distal TRAV genes. We confirmed the in vivo rescue of B-cell development with normal immunoglobulin M surface expression and a significant decrease in CD56bright natural killer cells. Together, we provide specificity, toxicity, and efficacy data supporting the development of a gene-correction therapy to benefit RAG2-deficient patients.

Generation of the transgene-free canker-resistant Citrus sinensis using Cas12a/crRNA ribonucleoprotein in the T0 generation.

Citrus canker caused by Xanthomonas citri subsp. citri (Xcc) is a destructive citrus disease worldwide. Generating disease-resistant cultivars is the most effective, environmentally friendly and economic approach for disease control. However, citrus traditional breeding is lengthy and laborious. Here, we develop transgene-free canker-resistant Citrus sinensis lines in the T0 generation within 10 months through transformation of embryogenic protoplasts with Cas12a/crRNA ribonucleoprotein to edit the canker susceptibility gene CsLOB1. Among the 39 regenerated lines, 38 are biallelic/homozygous mutants, demonstrating a 97.4% biallelic/homozygous mutation rate. No off-target mutations are detected in the edited lines. Canker resistance of the cslob1-edited lines results from both abolishing canker symptoms and inhibiting Xcc growth. The transgene-free canker-resistant C. sinensis lines have received regulatory approval by USDA APHIS and are exempted from EPA regulation. This study provides a sustainable and efficient citrus canker control solution and presents an efficient transgene-free genome-editing strategy for citrus and other crops.

Propensity of Patient-Derived iPSCs for Retinal Differentiation: Implications for Autologous Cell Replacement.

Prior to use, newly generated induced pluripotent stem cells (iPSC) should be thoroughly validated. While excellent validation and release testing assays designed to evaluate potency, genetic integrity, and sterility exist, they do not have the ability to predict cell type-specific differentiation capacity. Selection of iPSC lines that have limited capacity to produce high-quality transplantable cells, places significant strain on valuable clinical manufacturing resources. The purpose of this study was to determine the degree and root cause of variability in retinal differentiation capacity between cGMP-derived patient iPSC lines. In turn, our goal was to develop a release testing assay that could be used to augment the widely used ScoreCard panel. IPSCs were generated from 15 patients (14-76 years old), differentiated into retinal organoids, and scored based on their retinal differentiation capacity. Despite significant differences in retinal differentiation propensity, RNA-sequencing revealed remarkable similarity between patient-derived iPSC lines prior to differentiation. At 7 days of differentiation, significant differences in gene expression could be detected. Ingenuity pathway analysis revealed perturbations in pathways associated with pluripotency and early cell fate commitment. For example, good and poor producers had noticeably different expressions of OCT4 and SOX2 effector genes. QPCR assays targeting genes identified via RNA sequencing were developed and validated in a masked fashion using iPSCs from 8 independent patients. A subset of 14 genes, which include the retinal cell fate markers RAX, LHX2, VSX2, and SIX6 (all elevated in the good producers), were found to be predictive of retinal differentiation propensity.

Boosting genome editing efficiency in human cells and plants with novel LbCas12a variants.

BACKGROUND: Cas12a (formerly known as Cpf1), the class II type V CRISPR nuclease, has been widely used for genome editing in mammalian cells and plants due to its distinct characteristics from Cas9. Despite being one of the most robust Cas12a nucleases, LbCas12a in general is less efficient than SpCas9 for genome editing in human cells, animals, and plants. RESULTS: To improve the editing efficiency of LbCas12a, we conduct saturation mutagenesis in E. coli and identify 1977 positive point mutations of LbCas12a. We selectively assess the editing efficiency of 56 LbCas12a variants in human cells, identifying an optimal LbCas12a variant (RVQ: G146R/R182V/E795Q) with the most robust editing activity. We further test LbCas12a-RV, LbCas12a-RRV, and LbCas12a-RVQ in plants and find LbCas12a-RV has robust editing activity in rice and tomato protoplasts. Interestingly, LbCas12a-RRV, resulting from the stacking of RV and D156R, displays improved editing efficiency in stably transformed rice and poplar plants, leading to up to 100% editing efficiency in T0 plants of both plant species. Moreover, this high-efficiency editing occurs even at the non-canonical TTV PAM sites. CONCLUSIONS: Our results demonstrate that LbCas12a-RVQ is a powerful tool for genome editing in human cells while LbCas12a-RRV confers robust genome editing in plants. Our study reveals the tremendous potential of these LbCas12a variants for advancing precision genome editing applications across a wide range of organisms.

Comparative analysis of CRISPR off-target discovery tools following ex vivo editing of CD34+ hematopoietic stem and progenitor cells.

While a number of methods exist to investigate CRISPR off-target (OT) editing, few have been compared head-to-head in primary cells after clinically relevant editing processes. Therefore, we compared in silico tools (COSMID, CCTop, and Cas-OFFinder) and empirical methods (CHANGE-Seq, CIRCLE-Seq, DISCOVER-Seq, GUIDE-Seq, and SITE-Seq) after ex vivo hematopoietic stem and progenitor cell (HSPC) editing. We performed editing using 11 different gRNAs complexed with Cas9 protein (high-fidelity [HiFi] or wild-type versions), then performed targeted next-generation sequencing of nominated OT sites identified by in silico and empirical methods. We identified an average of less than one OT site per guide RNA (gRNA) and all OT sites generated using HiFi Cas9 and a 20-nt gRNA were identified by all OT detection methods with the exception of SITE-seq. This resulted in high sensitivity for the majority of OT nomination tools and COSMID, DISCOVER-Seq, and GUIDE-Seq attained the highest positive predictive value (PPV). We found that empirical methods did not identify OT sites that were not also identified by bioinformatic methods. This study supports that refined bioinformatic algorithms could be developed that maintain both high sensitivity and PPV, thereby enabling more efficient identification of potential OT sites without compromising a thorough examination for any given gRNA.

Drag-and-drop genome insertion of large sequences without double-strand DNA cleavage using CRISPR-directed integrases.

Programmable genome integration of large, diverse DNA cargo without DNA repair of exposed DNA double-strand breaks remains an unsolved challenge in genome editing. We present programmable addition via site-specific targeting elements (PASTE), which uses a CRISPR-Cas9 nickase fused to both a reverse transcriptase and serine integrase for targeted genomic recruitment and integration of desired payloads. We demonstrate integration of sequences as large as ~36 kilobases at multiple genomic loci across three human cell lines, primary T cells and non-dividing primary human hepatocytes. To augment PASTE, we discovered 25,614 serine integrases and cognate attachment sites from metagenomes and engineered orthologs with higher activity and shorter recognition sequences for efficient programmable integration. PASTE has editing efficiencies similar to or exceeding those of homology-directed repair and non-homologous end joining-based methods, with activity in non-dividing cells and in vivo with fewer detectable off-target events. PASTE expands the capabilities of genome editing by allowing large, multiplexed gene insertion without reliance on DNA repair pathways.

Amount of Cas9 protein introduced into mouse embryos via electroporation affects the genome-editing rate.

Genetically engineered animals can be produced quickly using genome editing technology. A new electroporation technique, technique for animal knockout system by electroporation (TAKE), aids in the production of genome-edited animals by introducing nucleases into intact embryos using electroporation instead of microinjection. It is difficult to confirm nuclease delivery into embryos after electroporation using the conventional TAKE method. We previously reported the successful visualization of fluorescently-labeled tracrRNA in embryos after electroporation Cas9 paired with the crRNA:tracrRNA-ATTO550 duplex. However, the amount of fluorescence signal from labeled tracrRNA in embryos did not correlate with the genome editing rate of the offspring. This study examined the visualization of Cas9 protein in embryos after electroporation and its correlation with the genome editing rate of the offspring using a fluorescent Cas9 fusion protein. The fluorescent Cas9 protein was observed in all embryos that survived following electroporation. We found that the efficiency of Cas9 protein delivery into embryos via electroporation depended on the pulse length. Furthermore, we demonstrated that the amount of fluorescent Cas9 protein detected in the embryos correlated with the genome editing efficiency of the embryos. These data indicate that the TAKE method using fluorescently-labeled nucleases can be used to optimize the delivery conditions and verify nuclease delivery into individual embryos prior to embryo transfer for the efficient production of genome-edited animals.

Highly Efficient Genome Editing in Plant Protoplasts by Ribonucleoprotein Delivery of CRISPR-Cas12a Nucleases.

Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) mediated genome editing is a powerful approach for crop improvement. Traditional transformation methods based on plasmid delivery pose concerns associated with transgene integration and off-target effects. CRISPR delivered as ribonucleoproteins (RNPs) can prevent exogenous DNA integration, minimize off-target effects, and reduce cellular toxicity. Although RNP delivered CRISPR genome editing has been demonstrated in many plant species, optimization strategies that yield high editing efficiencies have not been thoroughly investigated. Using rice and citrus protoplast systems we demonstrated highly efficient genome editing using Cas12a delivered as RNPs. Four Cas12a variants, including LbCas12a, LbCas12a-E795L, AsCas12a, and AsCas12a Ultra, were investigated. Nearly 100% editing efficiency was observed for three out of four target sites by LbCas12a, LbCas12a-E795L, and AsCas12a Ultra, as measured by restriction fragment length polymorphism (RFLP) and verified by next generation sequencing of PCR amplicons. RNP delivery resulted in higher editing efficiencies than plasmid delivery at 32°C and 25°C. LbCas12a and LbCas12a-E795L demonstrated increased editing efficiencies in comparison to AsCas12a and AsCas12a Ultra, especially when used at lower RNP concentrations. In addition, we discovered that a 1:1 Cas12a:crRNA molar ratio is sufficient to achieve efficient genome editing. Nuclear localization signals (NLSs) are essential for efficient RNP-based genome editing. However, the different crRNA modifications tested did not significantly improve genome editing efficiency. Finally, we applied the Cas12a RNP system in citrus protoplasts and obtained similarly high editing efficiencies at the target site. Our study provides a comprehensive guideline for Cas12a-mediated genome editing using RNP delivery in plant cells, setting the foundation for the generation of transgene-free genome edited plants.

Efficient Targeted Mutagenesis Mediated by CRISPR-Cas12a Ribonucleoprotein Complexes in Maize.

Recent advances in the development of CRISPR-Cas genome editing technologies have made it possible to perform targeted mutagenesis and precise gene replacement in crop plants. CRISPR-Cas9 and CRISPR-Cas12a are two main types of widely used genome editing systems. However, when CRISPR-Cas12a editing machinery is expressed from a transgene, some chromosomal targets encountered low editing frequency in important crops like maize and soybean. Here, we report efficient methods to directly generate genome edited lines by delivering Cas12a-gRNA ribonucleoprotein complex (RNP) to immature maize embryos through particle bombardment in an elite maize variety. Genome edited lines were obtained at ~7% frequency without any selection during regeneration via biolistic delivery of Cas12a RNP into immature embryos. Strikingly, the gene editing rate was increased to 60% on average and up to 100% in some experiments when the Cas12a RNP was co-delivered with a PMI selectable marker gene cassette and the induced callus cultures were selected with mannose. We also show that use of higher activity Cas12a mutants resulted in improved editing efficiency in more recalcitrant target sequence. The advances described here provide useful tools for genetic improvement of maize.

Development of ß-globin gene correction in human hematopoietic stem cells as a potential durable treatment for sickle cell disease.

Sickle cell disease (SCD) is the most common serious monogenic disease with 300,000 births annually worldwide. SCD is an autosomal recessive disease resulting from a single point mutation in codon six of the ß-globin gene (HBB). Ex vivo ß-globin gene correction in autologous patient-derived hematopoietic stem and progenitor cells (HSPCs) may potentially provide a curative treatment for SCD. We previously developed a CRISPR-Cas9 gene targeting strategy that uses high-fidelity Cas9 precomplexed with chemically modified guide RNAs to induce recombinant adeno-associated virus serotype 6 (rAAV6)-mediated HBB gene correction of the SCD-causing mutation in HSPCs. Here, we demonstrate the preclinical feasibility, efficacy, and toxicology of HBB gene correction in plerixafor-mobilized CD34+ cells from healthy and SCD patient donors (gcHBB-SCD). We achieved up to 60% HBB allelic correction in clinical-scale gcHBB-SCD manufacturing. After transplant into immunodeficient NSG mice, 20% gene correction was achieved with multilineage engraftment. The long-term safety, tumorigenicity, and toxicology study demonstrated no evidence of abnormal hematopoiesis, genotoxicity, or tumorigenicity from the engrafted gcHBB-SCD drug product. Together, these preclinical data support the safety, efficacy, and reproducibility of this gene correction strategy for initiation of a phase 1/2 clinical trial in patients with SCD.

AsCas12a ultra nuclease facilitates the rapid generation of therapeutic cell medicines.

Though AsCas12a fills a crucial gap in the current genome editing toolbox, it exhibits relatively poor editing efficiency, restricting its overall utility. Here we isolate an engineered variant, "AsCas12a Ultra", that increased editing efficiency to nearly 100% at all sites examined in HSPCs, iPSCs, T cells, and NK cells. We show that AsCas12a Ultra maintains high on-target specificity thereby mitigating the risk for off-target editing and making it ideal for complex therapeutic genome editing applications. We achieved simultaneous targeting of three clinically relevant genes in T cells at >90% efficiency and demonstrated transgene knock-in efficiencies of up to 60%. We demonstrate site-specific knock-in of a CAR in NK cells, which afforded enhanced anti-tumor NK cell recognition, potentially enabling the next generation of allogeneic cell-based therapies in oncology. AsCas12a Ultra is an advanced CRISPR nuclease with significant advantages in basic research and in the production of gene edited cell medicines.

Gene replacement of α-globin with ß-globin restores hemoglobin balance in ß-thalassemia-derived hematopoietic stem and progenitor cells.

ß-Thalassemia pathology is due not only to loss of ß-globin (HBB), but also to erythrotoxic accumulation and aggregation of the ß-globin-binding partner, α-globin (HBA1/2). Here we describe a Cas9/AAV6-mediated genome editing strategy that can replace the entire HBA1 gene with a full-length HBB transgene in ß-thalassemia-derived hematopoietic stem and progenitor cells (HSPCs), which is sufficient to normalize ß-globin:α-globin messenger RNA and protein ratios and restore functional adult hemoglobin tetramers in patient-derived red blood cells. Edited HSPCs were capable of long-term and bilineage hematopoietic reconstitution in mice, establishing proof of concept for replacement of HBA1 with HBB as a novel therapeutic strategy for curing ß-thalassemia.

Efficiency, Specificity and Temperature Sensitivity of Cas9 and Cas12a RNPs for DNA-free Genome Editing in Plants.

Delivery of genome editing reagents using CRISPR-Cas ribonucleoproteins (RNPs) transfection offers several advantages over plasmid DNA-based delivery methods, including reduced off-target editing effects, mitigation of random integration of non-native DNA fragments, independence of vector constructions, and less regulatory restrictions. Compared to the use in animal systems, RNP-mediated genome editing is still at the early development stage in plants. In this study, we established an efficient and simplified protoplast-based genome editing platform for CRISPR-Cas RNP delivery, and then evaluated the efficiency, specificity, and temperature sensitivity of six Cas9 and Cas12a proteins. Our results demonstrated that Cas9 and Cas12a RNP delivery resulted in genome editing frequencies (8.7-41.2%) at various temperature conditions, 22°C, 26°C, and 37°C, with no significant temperature sensitivity. LbCas12a often exhibited the highest activities, while AsCas12a demonstrated higher sequence specificity. The high activities of CRISPR-Cas RNPs at 22° and 26°C, the temperature preferred by plant transformation and tissue culture, led to high mutagenesis efficiencies (34.0-85.2%) in the protoplast-regenerated calli and plants with the heritable mutants recovered in the next generation. This RNP delivery approach was further extended to pennycress (Thlaspi arvense), soybean (Glycine max) and Setaria viridis with up to 70.2% mutagenesis frequency. Together, this study sheds light on the choice of RNP reagents to achieve efficient transgene-free genome editing in plants.

Systematic in vitro profiling of off-target affinity, cleavage and efficiency for CRISPR enzymes.

CRISPR RNA-guided endonucleases (RGEs) cut or direct activities to specific genomic loci, yet each has off-target activities that are often unpredictable. We developed a pair of simple in vitro assays to systematically measure the DNA-binding specificity (Spec-seq), catalytic activity specificity (SEAM-seq) and cleavage efficiency of RGEs. By separately quantifying binding and cleavage specificity, Spec/SEAM-seq provides detailed mechanistic insight into off-target activity. Feature-based models generated from Spec/SEAM-seq data for SpCas9 were consistent with previous reports of its in vitro and in vivo specificity, validating the approach. Spec/SEAM-seq is also useful for profiling less-well characterized RGEs. Application to an engineered SpCas9, HiFi-SpCas9, indicated that its enhanced target discrimination can be attributed to cleavage rather than binding specificity. The ortholog ScCas9, on the other hand, derives specificity from binding to an extended PAM. The decreased off-target activity of AsCas12a (Cpf1) appears to be primarily driven by DNA-binding specificity. Finally, we performed the first characterization of CasX specificity, revealing an all-or-nothing mechanism where mismatches can be bound, but not cleaved. Together, these applications establish Spec/SEAM-seq as an accessible method to rapidly and reliably evaluate the specificity of RGEs, Cas::gRNA pairs, and gain insight into the mechanism and thermodynamics of target discrimination.

Evaluation and Reduction of CRISPR Off-Target Cleavage Events.

Introduction of CRISPR/Cas9 methods (clustered regularly interspaced short palindromic repeats, CRISPR-associated protein 9) have led to a huge surge in the use of precision genome editing for research applications. Translational medical efforts are likewise rapidly progressing, and Phase I clinical trials using these techniques have already started. As with any new technology that is applied to medical therapeutics, risks must be carefully defined and steps taken to mitigate side effects wherever possible. Effective methods are now available that permit identification of off-target cleavage events, a major class of potential side effects seen in mammalian genome editing. Off-target prediction algorithms are improving and have utility, but are insufficient to use alone. Empiric methods to define the off-target profile must also be used. Once defined, the frequency of off-target cleavage can be minimized using methods that limit the duration of exposure of the genome to the active genome editing complex, for example, using the ribonucleoprotein (RNP) approach. In addition, Cas9 mutants have been developed that markedly reduce the rate of off-target cleavage compared to the wild-type enzyme. Use of these new tools should become standard practice for medical applications.

Identification of preexisting adaptive immunity to Cas9 proteins in humans.

The CRISPR-Cas9 system is a powerful tool for genome editing, which allows the precise modification of specific DNA sequences. Many efforts are underway to use the CRISPR-Cas9 system to therapeutically correct human genetic diseases1-6. The most widely used orthologs of Cas9 are derived from Staphylococcus aureus and Streptococcus pyogenes5,7. Given that these two bacterial species infect the human population at high frequencies8,9, we hypothesized that humans may harbor preexisting adaptive immune responses to the Cas9 orthologs derived from these bacterial species, SaCas9 (S. aureus) and SpCas9 (S. pyogenes). By probing human serum for the presence of anti-Cas9 antibodies using an enzyme-linked immunosorbent assay, we detected antibodies against both SaCas9 and SpCas9 in 78% and 58% of donors, respectively. We also found anti-SaCas9 T cells in 78% and anti-SpCas9 T cells in 67% of donors, which demonstrates a high prevalence of antigen-specific T cells against both orthologs. We confirmed that these T cells were Cas9-specific by demonstrating a Cas9-specific cytokine response following isolation, expansion, and antigen restimulation. Together, these data demonstrate that there are preexisting humoral and cell-mediated adaptive immune responses to Cas9 in humans, a finding that should be taken into account as the CRISPR-Cas9 system moves toward clinical trials.

A high-fidelity Cas9 mutant delivered as a ribonucleoprotein complex enables efficient gene editing in human hematopoietic stem and progenitor cells.

Translation of the CRISPR-Cas9 system to human therapeutics holds high promise. However, specificity remains a concern especially when modifying stem cell populations. We show that existing rationally engineered Cas9 high-fidelity variants have reduced on-target activity when using the therapeutically relevant ribonucleoprotein (RNP) delivery method. Therefore, we devised an unbiased bacterial screen to isolate variants that retain activity in the RNP format. Introduction of a single point mutation, p.R691A, in Cas9 (high-fidelity (HiFi) Cas9) retained the high on-target activity of Cas9 while reducing off-target editing. HiFi Cas9 induces robust AAV6-mediated gene targeting at five therapeutically relevant loci (HBB, IL2RG, CCR5, HEXB, and TRAC) in human CD34+ hematopoietic stem and progenitor cells (HSPCs) as well as primary T cells. We also show that HiFi Cas9 mediates high-level correction of the sickle cell disease (SCD)-causing p.E6V mutation in HSPCs derived from patients with SCD. We anticipate that HiFi Cas9 will have wide utility for both basic science and therapeutic genome-editing applications.

Global role of the bacterial post-transcriptional regulator CsrA revealed by integrated transcriptomics.

CsrA is a post-transcriptional regulatory protein that is widely distributed among bacteria. This protein influences bacterial lifestyle decisions by binding to the 5' untranslated and/or early coding regions of mRNA targets, causing changes in translation initiation, RNA stability, and/or transcription elongation. Here, we assess the contribution of CsrA to gene expression in Escherichia coli on a global scale. UV crosslinking immunoprecipitation and sequencing (CLIP-seq) identify RNAs that interact directly with CsrA in vivo, while ribosome profiling and RNA-seq uncover the impact of CsrA on translation, RNA abundance, and RNA stability. This combination of approaches reveals unprecedented detail about the regulatory role of CsrA, including novel binding targets and physiological roles, such as in envelope function and iron homeostasis. Our findings highlight the integration of CsrA throughout the E. coli regulatory network, where it orchestrates vast effects on gene expression.