Unedited Cells Research Articles

O18 Precision genome editing for targeted correction of pathogenic D50N mutation in keratitis–ichthyosis–deafness syndrome using CRISPR/Cas9 and homology-directed repair

Abstract Introduction and aims Keratitis–ichthyosis–deafness (KID) syndrome is an ectodermal disorder that causes blindness, skin inflammation and deafness. It is caused by autosomal dominant mutations in the gap junction beta 2 (GJB2) gene with 86% being a hotspot mutation on c.148G>A, resulting in the amino acid replacement (D50N) in its coding protein connexin 26 (Cx26). There is currently no curative treatment for KID syndrome. We aim to correct the hotspot mutation D50N using a genome-editing approach. Methods Keratinocytes generated from the patient with KID who had a D50N mutation (KID-KCs) were genetically corrected by homologous-directed repair/CRISPR-Cas9 using an electroporation approach. The correction efficiency was determined and analysed using Sanger sequencing and Synthego Inference of CRISPR Edits analysis. The off-target effects were confirmed by whole-exome sequencing [exome next-generation sequencing (NGS)]. Furthermore, the functional recovery following genome editing was assessed by GJB2 mRNA production, Cx26 protein expression, and trafficking and hemichannel activity. Results The correction efficiencies ranged from 95% to 100% with different doses of CRISPR-Cas9 and 8µg CRISPR-Cas9 was the most optimal concentration with the efficiency of 100% and approximately 0% insertion-deletion (INDEL). Fifty days after gene editing, the INDEL increased to only 2%, indicating a minimum residual CRISPR-Cas9 effect in edited cells. The exome-NGS analysis also showed no off-target effects in the outside target area. Functional studies showed that there was a significant increase in wildtype GJB2 mRNA expression and a reduction of mutant GJB2 expression (n = 5, P < 0.01). Increased Cx26 membranous localization in KID-KCs was also observed (n = 3, P < 0.01). Additionally, Cx26 hemichannel activity assessment using the neurobiotin assay showed a significant decrease in ‘leaky’ hemichannel in gene-edited KID-KCs compared with unedited cells (n = 3, P < 0.05). Conclusions We can genome-correct KID keratinocytes harbouring the mutation D50N with 100% editing efficiency, 0–2% INDEL, and undetectable off-target effects. This result indicates a promising gene therapy for KID syndrome with the hotspot mutation D50N.

Expanding the flexibility of base editing for high-throughput genetic screens in bacteria.

Genome-wide screens have become powerful tools for elucidating genotype-to-phenotype relationships in bacteria. Of the varying techniques to achieve knockout and knockdown, CRISPR base editors are emerging as promising options. However, the limited number of available, efficient target sites hampers their use for high-throughput screening. Here, we make multiple advances to enable flexible base editing as part of high-throughput genetic screening in bacteria. We first co-opt the Streptococcus canis Cas9 that exhibits more flexible protospacer-adjacent motif recognition than the traditional Streptococcus pyogenes Cas9. We then expand beyond introducing premature stop codons by mutating start codons. Next, we derive guide design rules by applying machine learning to an essentiality screen conducted in Escherichia coli. Finally, we rescue poorly edited sites by combining base editing with Cas9-induced cleavage of unedited cells, thereby enriching for intended edits. The efficiency of this dual system was validated through a conditional essentiality screen based on growth in minimal media. Overall, expanding the scope of genome-wide knockout screens with base editors could further facilitate the investigation of new gene functions and interactions in bacteria.

Streamlined, single-step non-viral CRISPR-Cas9 knockout strategy enhances gene editing efficiency in primary human chondrocyte populations

BackgroundCRISPR-Cas9-based genome engineering represents a powerful therapeutic tool for cartilage tissue engineering and for understanding molecular pathways driving cartilage diseases. However, primary chondrocytes are difficult to transfect and rapidly dedifferentiate during monolayer (2D) cell culture, making the lengthy expansion of a single-cell-derived edited clonal population not feasible. For this reason, functional genetics studies focused on cartilage and rheumatic diseases have long been carried out in cellular models that poorly recapitulate the native molecular properties of human cartilaginous tissue (e.g., cell lines, induced pluripotent stem cells). Here, we set out to develop a non-viral CRISPR-Cas9, bulk-gene editing method suitable for chondrocyte populations from different cartilaginous sources.MethodsWe screened electroporation and lipid nanoparticles for ribonucleoprotein (RNP) delivery in primary polydactyly chondrocytes, and optimized RNP reagents assembly. We knocked out RELA (also known as p65), a subunit of the nuclear factor kappa B (NF-κB), in polydactyly chondrocytes and further characterized knockout (KO) cells with RT-qPCR and Western Blot. We tested RELA KO in chondrocytes from diverse cartilaginous sources and characterized their phenotype with RT-qPCR. We examined the chondrogenic potential of wild-type (WT) and KO cell pellets in presence and absence of interleukin-1β (IL-1β).ResultsWe established electroporation as the optimal transfection technique for chondrocytes enhancing transfection and editing efficiency, while preserving high cell viability. We knocked out RELA with an unprecedented efficiency of ~90%, confirming lower inflammatory pathways activation upon IL-1β stimulation compared to unedited cells. Our protocol could be easily transferred to primary human chondrocytes harvested from osteoarthritis (OA) patients, human FE002 chondroprogenitor cells, bovine chondrocytes, and a human chondrocyte cell line, achieving comparable mean RELA KO editing levels using the same protocol. All KO pellets from primary human chondrocytes retained chondrogenic ability equivalent to WT cells, and additionally displayed enhanced matrix retention under inflamed conditions.ConclusionsWe showcased the applicability of our bulk gene editing method to develop effective autologous and allogeneic off-the-shelf gene therapies strategies and to enable functional genetics studies in human chondrocytes to unravel molecular mechanisms of cartilage diseases.

Dual α-Globin and Truncated EPO Receptor Knockin Restores Hemoglobin Production in α-Thalassemia-Derived Hematopoietic Stem and Progenitor Cells

Introduction: α-thalassemia major (ATM) is an autosomal recessive disorder where all four copies of the α-globin gene are deleted. A traditionally lethal disease due to severe fetal hypoxia, this condition is now survivable due to the advent of in utero blood transfusions. However, patients still require life-long transfusions postnatally. While allogeneic hematopoietic stem cell transplant (HSCT) can provide a definitive cure, it is limited by the paucity of suitable donors, has a high morbidity and mortality due the prerequisite myeloablative regimens, and carries the risk of immune rejection and graft-versus-host disease. Here, we describe a CRISPR/AAV-mediated genome editing strategy to restore a full-length copy of the α-globin gene at the β-globin locus in ATM patient-derived HSCs to restore normal hemoglobin production to a patient's red blood cells (RBCs). Methods: We identified a Cas9 gRNA that efficiently introduced indels at the HBB locus as well as an AAV6 vector that mediated efficient integration into the start codon of HBB. After designing and testing integration cassettes, we developed two candidate vectors that insert a HBA1 transgene at the HBB locus, with one bicistronic vector that also expresses a truncated form of the erythropoietin receptor (t EPOR) to increase erythropoietic output from edited HSCs. We used this approach to edit CD34 + HSCs derived from the bone marrow of three patients with ATM. Edited cells were cultured for 16 days in RBC differentiation media. Editing rates were quantified by a custom ddPCR assay. To determine the transcriptional response to these editing strategies, bulk RNA sequencing was performed comparing wild-type cells and patient cells that underwent respective editing conditions. Lastly, α-globin protein production was quantified using tetramer and single-globin high protein liquid chromatography (HPLC). Results: While effective editing frequencies in ATM HSCs were achieved by the HBA integration vector (39% edited alleles), we observed that editing rates increased over the course of RBC differentiation in the combined HBA+t EPOR edited condition, reaching 76% edited alleles by day 16. We also found that total cell counts by the end of RBC differentiation more than doubled (2.2-fold) in the HBA+t EPOR condition compared to the HBA alone condition. RNA sequencing of edited ATM-derived HSCs following RBC differentiation revealed an upregulation of HBA1, HBA2, and EPOR transcripts compared to unedited patient cells. These results were supported by HPLC, which demonstrated restoration of hemoglobin tetramers, including fetal hemoglobin (α+γ subunits), adult hemoglobin (α+β subunits), and hemoglobin A2 (α+δ subunits) compared to unedited patient cells. Complementing this with globin single chain analysis found that we had achieved 11% restoration of α-globin compared to wild-type levels. In spite of integration at the HBB locus, neither RNA sequencing nor HPLC showed disruption of β-globin production. α-globin production with a concomitant decrease in the formation of toxic β-globin dimers was confirmed by Western blot, with restoration of the normal balance of α-globin to dimeric β-globin in edited cells to wild-type levels. Conclusions: To our knowledge, these results demonstrate for the first time that gene therapy or genome editing may be used to increase α-globin production in ATM patient-derived RBCs. We found that the editing strategy that paired site-specific α-globin integration with t EPOR expression achieved the highest editing frequencies, greatest production of edited cells, and the most robust production of α-globin protein compared to α-globin integration alone. By increasing erythropoietic output using t EPOR, this approach has the potential to overcome some of the clinical challenges previously seen in HSCT for the hemoglobinopathies, including low editing and engraftment rates. Moreover, as patients with severe anemia such those with ATM have increased levels of circulating erythropoietin due to severe anemia, a t EPOR-based enrichment strategy is likely to be especially potent following transplantation into ATM bone marrow. We believe this could allow clinicians to reduce or eliminate the need for myeloablation, which stand as a major barrier to HSCT safety and efficacy. These findings support development of a definitive ex vivo autologous genome editing strategy that may be curative for ATM.

A Pipeline for Comparison and Selection of Clinically Applicable Gene Therapy Approaches for ELANE-Associated Severe Congenital Neutropenia

Several gene editing strategies for autosomal dominant ELANE-associated severe congenital neutropenia (CN) have been proposed. All strategies efficiently correct CN with different degrees of the investigated comprehensiveness of toxicity and off-target activity. The next step now is to select the best approach for clinical translation. The first-line awareness should consider not only the universal applicability of gene editing but, most importantly, its safety. Since CN patients have a high cumulative incidence of leukemogenic transformation and pre-leukemia cells in the patient's bone marrow are detected at an early age, safety considerations are of the utmost importance. We compared gene-editing methods head-to-head in CD34 + cells of two patients with hot-spot ELANE mutations associated with a poor response to G-CSF and high risk for leukemia development, p.A57V and p.G214V. We edited cells with the universal ELANE knockout targeting wild-type and mutant alleles in exon 2 ( uEKO). We also repaired the mutated ELANE by inducing a double-stranded break (DSB) at the position of mutation with the delivery of the WT ELANE template for homology-driven repair (HDR) by AAV6 ( HDR-AAV). We also tested the allele-specific knockout of the mutated allele ( ASKO) and the allele-specific HDR approach that relies on ssODNs to deliver the wild-type sequence for p.G214V mutation ( AS-HDR). We investigated the effects of gene editing on the survival and granulocytic differentiation of gene-edited cells as well as editing efficiency and patient-specific genome-wide off-target profiles. Cell survival was between 74 % and 95 % in all groups post-gene editing. For patient 1 with p.A57V mutation, we observed gene editing of 62 % in the control-AAVS1 locus, 70 % in uEKO, 76 % in ASKO, and 10.3 % HDR with an additional 49.5 % of KO in HDR-AAV. The editing led to a significant increase in differentiated neutrophils from 14.5 % in the control group to 57.4 %, 55.2 %, and 45.3 % in uEKO, ASKO and HDR-AAV groups, respectively. Gene editing also facilitated a significant increase in CFU-G and CFU-GM colonies for all treatment conditions, as compared to the control. For patient 2 with p.G214V mutation, we measured 86 % indels in control-AAVS1, 34.4 % in uEKO and 80.25 % in HDR-AAV. In the AS-HDR group, no HDR was observed, but 92 % of mutant alleles contained frameshift indels. The edits resulted in a significant increase in neutrophils in liquid culture differentiation from 13.6 % in the control AAVS1group to 53.7 %, 57.3 %, and 62.8 % in uEKO, HDR-AAV, and AS-HDR, respectively. CFU-G and CFU-GM colonies were also enhanced. To confirm if allele-specific frameshifts in exon 5 restored granulopoiesis, we utilized a patient-derived iPSC cell line with the p.G214R mutation. We applied ASKO gene editing to this iPSC line and achieved 24 % ASKO. The editing improved neutrophil differentiation significantly, from 2.4 % in unedited cells to 25.55 % in edited cells. CFU assays also showed a significant increase in CFU-G and CFU-GM numbers. To detect off-targets on the patient genotype, we established GUIDE-Seq in patient-derived iPSCs with mutations p.A57V (iCN1) and p.G214R (iCN2). In iCN1, we found 10 off-targets (OT) for the uEKO sgRNA and 3 OTs for the ASKOsgRNA. For iCN2, we found 3 OTs for the HDR-AAV and the AS-HDR guides. These findings show that allele-specific guides have better OT profiles than guides targeting both alleles. Taken together, all tested approaches have low toxicity and comparable efficacy in restoring granulopoiesis in vitro. We found that editing above 30 % efficiently restored granulopoiesis, providing evidence for the therapeutic threshold. This highlights that safety and genome toxicity considerations should guide further clinical translation. To enable clinically relevant off-target activity detection of gene-editing in the patient's genome, GUIDE-Seq in patient-derived iPSCs should be performed. Further, our observation that the introduction of allele-specific frameshift indels in exon 5 can inactivate CN-causing mutations is unexpected and warrants further investigation. Head-to-head comparison of the in vivo engraftment of CD34+ cells gene-edited using several approaches should also be performed in the future.

Functional analysis of a putative HER2-associated expressed enhancer, Her2-Enhancer1, in breast cancer cells

HER-2/neu (HER2) is a member of the epidermal growth factor receptors family, encoding a protein with tyrosine kinase activity. Following the gene amplification or increased HER2 transcription, carcinogenesis has been observed in some cancers. Genetic and epigenetic changes occurring in enhancer sequences can deeply affect the expression and transcriptional regulation of downstream genes, which can cause some physiological and pathological changes, including tumor progression. A therapeutic approach that directly targets the genomic sequence alterations is of high importance, with low side effects on healthy cells. Here, we employed the CRISPR/Cas9 method to genetically knockout an expressed putative enhancer (GH17J039694; we coined it as Her2-Enhancer1) located within the HER2 gene, 17q12: 39,694,339–39,697,219 (UCSC-hg38). We then investigated the potential regulatory effect of Her2-Enhancer1 on HER2 and HER2-interacting genes. To evaluate the cis and trans effects of Her2-Enhancer1, genetic manipulation of this region was performed in HER2-positive and -negative breast cancer cells. Our bioinformatics and real-time PCR data revealed that this putative enhancer region is indeed expressed, and acts as an expressed enhancer. Further functional analysis on edited and unedited cells revealed a significant alteration in the expression of HER2 variants, as well as some other target genes of HER2. Moreover, the apoptosis rate was considerably elevated within the edited cells. As we expected, Western blot analysis confirmed a reduction in protein levels of HER2, GRB7, the gene interacting with HER2, and P-AKT in the PI3K/AKT pathway. Altogether, our findings revealed an enhancer regulatory role for Her2-Enhancer1 on HER2 and HER2-interacting genes; and that this region has a potential for targeted therapy of HER2-positive cancers.

Multiplex Genome Editing of Human T Cells with Innovative Transformer Base Editor (tBE) for Construction of Next Generation CAR-T Therapies

Autologous chimeric antigen receptor T (CAR-T) cell therapy has achieved great success for antitumor treatments especially against hematological malignancies, and several autologous CAR-T therapies have been approved around the world. However, high manufacturing cost and long manufacturing process hinder the broader application of autologous CAR-T cell therapy. As a result, off-the-shelf universal CAR-T cell therapy with donor derived healthy T cells is of great interest and under development by many companies globally. Gene editing technologies including TALENs, CRISPR/Cas nucleases and base editors are of great potential in T cell engineering for allogeneic T cell therapy. The innovative transformer base editor (tBE) is a base editing system that avoids to cause DNA double strand breaks (DSBs) and exhibits higher editing efficiency, no detected off-target mutations and lower cytotoxicity. Due to its capability to avoid off-target mutations and DSB-caused chromosomal translocations, tBE is potentially one of the best gene editing tools to induce multiplex editing in cells. Here we show that the tBE was used to induce highly efficient single or multiplex gene editing in primary human T cells. With optimized electroporation conditions for tBE, TRAC, CD52 and PDCD1 could be knocked out simultaneously with editing efficiency up to ~90% for each target gene. tBE triggered no detected off-target mutation or chromosomal abnormalities, which were reported in the cases of CRISPR/Cas- or TALEN-mediated multiplex editing. CD19-targeted CAR-T cells with triplex base editing exhibits enhanced antitumor efficacy in vitro and in mouse models compared with unedited CAR-T cells. Our results demonstrated the application of tBE for construction of next generation universal or enhanced CAR-T therapies.

Explore the alterations of downstream molecular pathways caused by ARID1A mutation/knockout in human endometrial cancer cells.

As one of the most common gynecologic malignancies, endometrial cancer (EC) is driven by multiple genetic alterations that may be targeted for treatments. AT-rich interaction domain 1A (ARID1A) gene mutations were reported as early events in endometrial carcinogenesis. To explore the alterations of downstream molecular pathways caused by ARID1A mutations and the associated therapeutic implications, we edited ARID1A gene in human endometrial cancer cell line Ishikawa using the Clustered Regularly Interspaced Short Palindromic Repeats and CRISPR-Associated Proteins (CRISPR/Cas9) technology. We successfully constructed a stable Ishikawa cell line with a confirmed 10bp deletion on the ARID1A gene, which resulted in a code-shift mutation and gene knockout. Compared with unedited wild-type cells, ARID1A knockout (KO) led to reduced apoptosis, accelerated transformation from G0/G1 to S phase, and enhanced cell proliferation. ARID1A deficiency would reduce the protein levels of p21, caspase 7, and caspase 9 in Ishikawa endometrial cancer cells compared with the wild-type cells. In addition, ARID1A KO resulted in high levels of microsatellite instability (MSI-H). Moreover, transcriptomic analyses showed that ARID1A KO can lead to activated phosphoinositide 3-kinase (PI3K)/protein kinase B (Akt) signaling. Furthermore, experimental analyses demonstrated that ARID1A KO cells had reduced expression of genetic instability-associated markers mutL homologue 1 (MLH1) and progesterone receptor B (PR) and increased p-Akt expression. These findings support further exploration of ARID1A as a therapeutic target for EC and provide insight into developing more effective treatments in EC, such as the combinatory use of immune checkpoint inhibitors.

Systematically attenuating DNA targeting enables CRISPR-driven editing in bacteria

Bacterial genome editing commonly relies on chromosomal cleavage with Cas nucleases to counter-select against unedited cells. However, editing normally requires efficient recombination and high transformation efficiencies, which are unavailable in most strains. Here, we show that systematically attenuating DNA targeting activity enables RecA-mediated repair in different bacteria, allowing chromosomal cleavage to drive genome editing. Attenuation can be achieved by altering the format or expression strength of guide (g)RNAs; using nucleases with reduced cleavage activity; or engineering attenuated gRNAs (atgRNAs) with disruptive hairpins, perturbed nuclease-binding scaffolds, non-canonical PAMs, or guide mismatches. These modifications greatly increase cell counts and even improve the efficiency of different types of edits for Cas9 and Cas12a in Escherichia coli and Klebsiella oxytoca. We further apply atgRNAs to restore ampicillin sensitivity in Klebsiella pneumoniae, establishing a resistance marker for genetic studies. Attenuating DNA targeting thus offers a counterintuitive means to achieve CRISPR-driven editing across bacteria.

A Missense Variant in CASKIN1's Proline-Rich Region Segregates with Psychosis in a Three-Generation Family.

The polygenic nature of schizophrenia (SCZ) implicates many variants in disease development. Rare variants of high penetrance have been shown to contribute to the disease prevalence. Whole-exome sequencing of a large three-generation family with SCZ and bipolar disorder identified a single segregating novel, rare, non-synonymous variant in the gene CASKIN1. The variant D1204N is absent from all databases, and CASKIN1 has a gnomAD missense score Z = 1.79 and pLI = 1, indicating its strong intolerance to variation. We find that introducing variants in the proline-rich region where the D1204N resides results in significant cellular changes in iPSC-derived neurons, consistent with CASKIN1’s known functions. We observe significant transcriptomic changes in 368 genes (padj < 0.05) involved in neuronal differentiation and nervous system development. We also observed nominally significant changes in the frequency of action potentials during differentiation, where the speed at which the edited and unedited cells reach the same level of activity differs. Our results suggest that CASKIN1 is an excellent gene candidate for psychosis development with high penetrance in this family.

Secreted PGK1 and IGFBP2 contribute to the bystander effect of miR-10b gene editing in glioma

MicroRNA-10b (miR-10b) is an essential glioma driver and one of the top candidates for targeted therapies for glioblastoma and other cancers. This unique miRNA controls glioma cell cycle and viability via an array of established conventional and unconventional mechanisms. Previously reported CRISPR-Cas9-mediated miR-10b gene editing of glioma cells invitro and established orthotopic glioblastoma in mouse models demonstrated the efficacy of this approach and its promise for therapy development. However, therapeutic gene editing in patients' brain tumors may be hampered, among other factors, by the imperfect delivery and distribution of targeting vectors. Here, we demonstrate that miR-10b gene editing in glioma cells triggers a potent bystander effect that leads to the selective cell death of the unedited glioma cells without affecting the normal neuroglial cells. The effect is mediated by the secreted miR-10b targets phosphoglycerate kinase 1 (PGK1) and insulin-like growth factor binding protein 2 (IGFBP2) that block cell-cycle progression and induce glioma cell death. These findings further support the feasibility of therapeutic miR-10b editing without the need to target every cell of the tumor.

Base Editing for Therapeutic Induction of Fetal Hemoglobin in β-Thalassemia

β-thalassemia, one of the most common inherited genetic disorders worldwide, is caused by more than 300 different mutations in the β-globin locus (HBB), all of which result in quantitative deficiency of β-globin protein with severe anemia and reduced or absent HbA (α2β2). Additionally, free α-globin protein forms toxic intracellular inclusions resulting in hemolysis of mature red blood cells, maturation arrest and apoptosis of erythroid precursors. The only approved cure for severe β-thalassemia is allogeneic hematopoietic stem cell transplantation. However, many patients lack ideal histocompatible donors and immune complications such as graft rejection and graft-versus-host disease can occur. These problems can be avoided by "gene therapies” in which the patient's own hematopoietic stem cells (HSCs) are isolated, genetically modified ex vivo and reinfused after administration of bone marrow conditioning to facilitate engraftment of the modified cells. Most β-thalassemia mutations can be treated by genetic manipulations that de-repress fetal-expressed γ-globin genes (HBG1 and HBG2) to induce fetal hemoglobin production (HbF, α2γ2) postnatally. Thus, Cas9 disruption of an erythroid enhancer in the γ-globin repressor gene BCL11A shows promising results in an ongoing clinical trial. However, multiple other strategies to induce HbF exist and the best one is not known. Compared to conventional Cas9 nuclease, base editors offer theoretical advantages, including the ability to create precise base pair substitutions without requiring genotoxic double strand DNA breaks. We used the adenine base editor ABE8e to generate the naturally occurring γ-globin -175 A>G variant, which causes hereditary persistence of fetal hemoglobin (HPFH), a benign genetic condition that alleviates co-inherited β-thalassemia. We edited human CD34+ hematopoietic stem and progenitor cells (HSPCs) from two individuals with severe transfusion dependent β-thalassemia by electroporating ribonucleoprotein (RNP) complexes of ABE8e-NG protein + single guide RNA (sgRNA) designed to install the γ-globin -175 A>G variant, followed by in vitro erythroid differentiation. On-target editing rates were 48-53% for donor 1 (n=3 replicates) and 22.9% for donor 2 (n=1). Analysis of burst forming unit erythroid (BFU-E) colonies revealed at least one on-target edit in 78% of donor 1 HSPCs and 63% of those from donor 2. Reverse phase HPLC of erythroid progeny showed increased β-like/α globin protein ratios: 0.92 ± 0.04 vs 0.53 ± 0.03 in unedited controls (p<0.0001) for donor 1, and 0.8 vs 0.5 in unedited controls for donor 2. Quantitative RT-PCR showed 6-8-fold increased γ-globin mRNA in the ABE-treated β-thalassemia cells. Ion exchange HPLC analysis of erythroid cell lysates showed reductions in free α-globin levels after ABE treatment: from 21.5 ± 2% of total globin protein in unedited cells to undetectable levels after editing in donor 1 and from 18% to 2.5% for donor 2. Maturation to the reticulocyte stage was also increased in base edited cells: 14.6 ± 2% vs 3.3 ± 1% in unedited controls for donor 1 (P<0.0001); and 26% vs 6.9% in unedited controls for donor 2. Reduction in apoptosis (37.4 ± 2.2% vs 23.6 ± 1.5% in unedited control cells, p=0.0003) and reactive oxygen species measured by DCFDA (MFI: 690 ± 97 vs 266 ± 9 in unedited control cells, p=0.009) was also seen in the ABE edited groups. To study editing outcomes in vivo, CD34+ HSPCs from donor 2 were treated with RNP and transplanted into immunodeficient NBSGW mice. After 16-18 weeks, human donor-derived cells were isolated from mouse recipient bone marrow and analyzed (n=4 mice per group). On-target editing frequencies in CD235a+ erythroblasts and CD34+ hematopoietic stem/progenitors were 17.5 ± 5% and 19.5 ±3%, respectively, compared to 22.9% in the input CD34+ cells. Similar to what we observed after in vitro differentiation, base edited erythroid cells generated in vivo exhibited improved globin chain balance with β-like/α globin protein ratios of 0.86 ± 0.12 vs 0.69 ± 0.04 in unedited controls, (p=0.04), reduced apoptosis (16.8 ± 4.2 % vs 26.6 ± 2.9 % in unedited control cells, p=0.004) and enhanced maturation into enucleated reticulocytes (14.4 ± 4.7 % vs 2.87 ± 1.4 % in unedited control cells, p=0.005). Our study provides proof of concept that ABE8e generation of the γ-globin -175 A>G HPFH variant in β-thalassemic HSCs represents a new therapeutic approach for β-thalassemia.

Increased Potency and Uniformity of Fetal Hemoglobin Induction from Base Editing Compared to Cas9 Nuclease

Genetic manipulations to increase fetal hemoglobin (HbF, α2γ2) in red blood cells (RBCs) can alleviate β-thalassemia and sickle cell disease (SCD). We and others demonstrated that CRIPSR/Cas9 disruption of γ-globin promoter binding motifs for the repressors BCL11A or ZBTB7A raises erythroid HbF expression to potentially therapeutic levels. Moreover, disruption of an erythroid BCL11A gene enhancer to induce HbF produced clinical improvement in humans with SCD. However, Cas9-induced double-stranded breaks (DSBs) can cause potentially deleterious chromosomal abnormalities. Base editing represents a promising approach to install precise nucleotide alterations through DSB-independent mechanisms. In contrast to Cas9 generated indels, base editors can recreate naturally occurring base pair substitutions that cause hereditary persistence of fetal hemoglobin (HPFH), including those that create de novo binding motifs for erythroid transcriptional activators at the γ-globin promoter. Base editing and Cas9 nuclease-based approaches to induce HbF have not been compared directly and the best therapeutic strategy is not known. We compared two Cas9-based strategies and three base editing strategies for HbF induction by modifying CD34+ hematopoietic stem and progenitor cells (HSPCs) and examining HbF expression in erythroid progeny generated by in vitro differentiation. We used Cas9 nuclease to disrupt the BCL11A gene erythroid enhancer or the BCL11A binding motif in the γ-globin promoter. Alternatively, we used the adenine base editor ABE7.10 to create base pair substitutions at γ-globin promoter positions -198, -175, and -113, to generate de-novo binding motifs for transcriptional activators KLF1, TAL1, and GATA1, respectively. The -175 A>G edit was most potent, producing an approximately 1% rise in HbF per % editing. In contrast, Cas9 disruption of the BCL11A enhancer or the BCL11A binding motif in the γ-globin promoter caused 0.33% and 0.30% increases in HbF per % editing, respectively. Cas9 nuclease but not base editing induced a TP53-mediated DNA damage response, as reflected by increased expression of CDKN1(P21) mRNA. To compare further the different edits, we analyzed burst forming unit-erythroid (BFU-E) colonies derived from modified CD34+ cells. Erythroid colonies with -175 A>G edit in all four γ-globin genes expressed 81±7% HbF compared to 14±11% in unedited controls. In contrast, HbF in BFU-E colonies was 33±19% after Cas9 disruption of the BCL11A repressor binding motif in the promoters of all γ-globin alleles and 50±14% after Cas9 disruption of the erythroid enhancer in the BCL11A gene. Among Cas9 mutations disrupting the γ-globin BCL11A binding motif, significantly increased HbF occurred with a 13 bp deletion but not with -1, -2, -3 or +1 indels. This finding is unexpected because elimination of BCL11A binding is believed to be sufficient for γ-globin induction. All high frequency indels targeting the BCL11A enhancer induced HbF similarly, albeit with greater variability than in colonies with the -175 A>G base edit. In normal donor CD34+ cells, ABE7.10 installed γ-globin -175 A>G at efficiencies of 43±6% (n=6). The newer, more active ABE8e produced a 1.4-fold improvement in editing efficiency and a 1.3-fold increase in HbF induction relative to ABE7.10. In preliminary studies, we achieved on-target editing of 62% with ABE8e in SCD patient CD34+ HSPCs. At 16 weeks after xenotransplantation of the modified HSPCs into NBSGW mice, on-target editing of donor-derived cells was 53±11%. Base edited erythroid cells isolated from recipient mouse bone marrow expressed 53±5% HbF, versus 2±1% in unedited cells. After xenotransplantation of the same donor CD34+ cells that were treated with Cas9 nuclease to disrupt the γ-globin promoter BCL11A binding motif, editing retention was 92±4%, with 29±12% HbF in donor derived erythroblasts. Thus, the -175 A>G edit produces superior HbF induction. Our data identifies a base editing strategy for potent, uniform induction of HbF and provides insights into γ-globin gene regulation by showing that disruption of the promoter BCL11A binding motif is not sufficient for transcriptional activation. More generally, we demonstrate that diverse indels generated by Cas9 nuclease cause unexpected variation in biological outcomes that can be circumvented by base editing, with important implications for therapeutic gene editing efforts.

Engineered Single Amino Acid Substitutions Protect Hematopoietic Stem and Progenitor Cells from CD123 Targeted Immunotherapy

Targeted eradication of transformed or otherwise dysregulated cells using monoclonal antibodies (mAb), T cell engagers (TCE) or chimeric antigen receptor (CAR) cells is very effective for hematologic diseases. CD123, the interleukin-3 (IL-3) receptor alpha-chain, is highly expressed in various hematological malignancies, including acute myeloid leukemia (AML) and blastic plasmacytoid dendritic cell neoplasm (BPDCN). However, shared expression of CD123 on hematopoietic stem and progenitor cells (HSPCs) bears the risk for extensive myelotoxicity upon targeted depletion. To enable therapeutic targeting of non-essential proteins post HSC transplantation (HSCT), it was proposed to engineer HSPCs that lack the target protein. Here, we show that expression and function of a genetically modified CD123 receptor can be preserved in genome engineered therapeutic cells despite full shielding from targeted therapy. We used the known co-crystal structure of CD123 bound to the mAb CSL362 (talacotuzumab) for computer-aided protein engineering. In silico, 28 candidate single amino acid (aa) substitutions at 3 key residues were designed to interfere with CSL362 binding while potentially preserving its function. Flow cytometry confirmed no binding of a CSL362 biosimilar to 13 variants and weak binding to 12 variants while 3 variants showed similar binding as wildtype (wt) CD123. Protein expression of all variants was comparable to wt CD123. Using real-time, label-free protein interaction measurements, we could not detect interactions between the non-binding variant receptors and the CSL362 biosimilar, even at high protein concentrations. Despite this, the non-binding variants preserved the normal binding of IL-3. Protein characterization of selected non- and weak binding variants demonstrated normal biophysical properties of all but one variant. All 13 non-binding variants abolished antibody dependent cellular cytotoxicity (ADCC) of the CSL362 IgG1 biosimilar in a surrogate reporter assay. In addition, a CSL362-derived TCE resulted in T cell activation and target cell killing of wt and strong binding CD123 variants while non-binding variants neither activated the T cells nor resulted in cell killing. Finally, the majority of non-binding variants (12/13) were completely protected from CSL362-CAR T cell cytotoxicity. To assess the function of the variant receptors we engineered the top 2 variants into the IL-3 dependent cell line TF-1 using CRISPR/Cas9-mediated homology directed repair (HDR). Engineered variant TF-1 cells demonstrated IL-3 dose-dependent growth comparable to their unedited counterparts. The addition of the CSL362 biosimilar inhibited IL-3 dependent cell growth in unedited TF-1 cells but not cells expressing variant receptors. Similarly, CRISPR/Cas9-engineered human CD34+ HSPC knock-in (KI) cells expressing the selected variants (Fig. 1a) were shielded from TCE killing using HLA-matched T cells in vitro (Fig. 1b). In addition, a set of in vitro experiments performed with CD34+ HSPC KI cells suggested normal differentiation potential and intact IL-3 signaling as measured by pSTAT5. In summary, our data demonstrate that single aa substitutions in CD123 introduced to HSPCs by CRISPR/Cas9-mediated HDR enable selective shielding from various immunotherapies, while preserving biophysical and functional properties. Variant IL-3 receptor engineered HSPC's and paired immunotherapies could enable novel and targeted therapeutic interventions post HSCT to either treat minimal residual disease or be used as a salvage therapy. Such approaches could expand the therapeutic window and broaden indications and applications for targeted immunotherapies for various malignant and benign conditions. Figure 1View largeDownload PPTFigure 1View largeDownload PPT Close modal

Efficient and Specific Multi-Locus Editing of Allogeneic CAR T Cells for Hypoimmunity during Large Scale Manufacture Using Cas12b

Introduction: Allogeneic CAR T cell therapies may offer key advantages over autologous therapies including choice of starting cells, better manufacturing control, and lower cost. However, the manufacture of these cells imposes a high demand on genome engineering specificity and efficiency, given the dual needs for product safety and yield. This challenge is amplified for newer generations of allogeneic CAR T cell designs that may offer improved performance but require more edits to produce, which may increase the potential of unintended genome modification and insufficient yield of fully engineered product. Here we have tested a novel editing platform, Cas12b, for the ability to mediate efficient and specific disruption of three genes (B2M, CIITA, and TRAC) during large scale manufacturing of CAR T cells with immune evasive properties and abrogated graft-vs-host response. We have characterized the resulting cell product for on-target modification efficiency, type and levels of unintended genome modification, and potency for tumor clearance in a Nalm6 humanized mouse model, with the aim of demonstrating suitability for translation to human studies. Methods: Our manufacturing strategy to generate hypoimmune CAR T cells introduces five genetic modifications, using Cas12b for disruption of B2M, CIITA, and TCR expression, and lentiviral transduction to overexpress CD47 and introduce a CD19 CAR. T cells obtained from healthy donors were first transduced with lentiviral vector and then edited simultaneously to disrupt the B2M, CIITA and TRAC genes via electroporation of Cas12b mRNA and appropriate guides. Edited and purified cells were characterized via flow cytometry for efficiency of each genetic modification. Cells were further characterized for knockout levels via next generation sequencing (NGS) as well as vector copy number per diploid genome (VCN). Cells were characterized for viability, proliferation, and efficacy in a Nalm6 mouse model of tumor clearance. An initial assessment of chromosomal translocations was performed by G banding karyotype analysis. To assess editing specificity, each Cas12b guide was evaluated by Cas-OFFinder and an unbiased oligonucleotide duplex capture assay in T cells from multiple donors under conditions of high on-target modification. Candidate off-target sites were then examined for evidence of cleavage via PCR amplification from the manufactured cell product followed by NGS. Results: Our manufacturing process yielded clinically relevant scales of cell product having > 85% B2M knockout, > 90% CIITA knockout, and < 1% of residual TCR expressing cells. VCN levels were < 5. Cell viability and proliferation were minimally impacted by the gene editing process. Cells exhibited robust anti-tumor properties with doses of 7 x 106 hypoimmune CD19 CAR T cells (per animal) providing efficient killing in a Nalm6 mouse model. Initial karyotype analysis identified low levels of translocation between on-target sites with overall translocation levels otherwise not significantly affected compared to unedited cells. Our characterization of Cas12b in edited T cells across multiple donors revealed high specificity, consistent with previous reports (Nat Commun 10 212), with studies to date having yielded no evidence of cleavage at any candidate off-target site. Summary: Cas12b-mediated multi-locus editing of allogeneic CAR T cells during large scale manufacture resulted in a high yield product with potent tumor clearance capabilities. Initial analysis of editing specificity across multiple donors and engineering runs identified no off-target sites, and early karyotype assessments indicated an overall level of chromosomal translocations consistent with prior experiences in the field. In companion in vivo studies, these hypoimmune CD19 CAR T cells have been shown to efficiently evade host immune response (see posters by Hu and Duback). Together, these data demonstrate a profile for potency, genome integrity, and manufacturing yield that supports progressing these cells into human studies.

A CRISPR-guided mutagenic DNA polymerase strategy for the detection of antibiotic-resistant mutations in M. tuberculosis

A sharp increase in multidrug-resistant tuberculosis (MDR-TB) threatens human health. Spontaneous mutation in essential gene confers an ability of Mycobacterium tuberculosis resistance to anti-TB drugs. However, conventional laboratory strategies for identification and prediction of the mutations in this slowly growing species remain challenging. Here, by combining XCas9 nickase and the error-prone DNA polymerase A from M. tuberculosis, we constructed a CRISPR-guided DNA polymerase system, CAMPER, for effective site-directed mutagenesis of drug-target genes in mycobacteria. CAMPER was able to generate mutagenesis of all nucleotides at user-defined loci, and its bidirectional mutagenesis at nick sites allowed editing windows with lengths up to 80 nucleotides. Mutagenesis of drug-targeted genes in Mycobacterium smegmatis and M. tuberculosis with this system significantly increased the fraction of the antibiotic-resistant bacterial population to a level approximately 60- to 120-fold higher than that in unedited cells. Moreover, this strategy could facilitate the discovery of the mutation conferring antibiotic resistance and enable a rapid verification of the growth phenotype-mutation genotype association. Our data demonstrate that CAMPER facilitates targeted mutagenesis of genomic loci and thus may be useful for broad functions such as resistance prediction and development of novel TB therapies.

Development of an efficient single-cell cloning and expansion strategy for genome edited induced pluripotent stem cells.

Disease-specific human induced pluripotent stem cells (hiPSCs) can be generated directly from individuals with known disease characteristics or alternatively be modified using genome editing approaches to introduce disease causing genetic mutations to study the biological response of those mutations. The genome editing procedure in hiPSCs is still inefficient, particularly when it comes to homology directed repair (HDR) of genetic mutations or targeted transgene insertion in the genome and single cell cloning of edited cells. In addition, genome editing processes also involve additional cellular stresses such as poor cell viability and genetic stability of hiPSCs. Therefore, efficient workflows are desired to increase genome editing application to hiPSC disease models and therapeutic applications. To this end, we demonstrate an efficient workflow for feeder-free single cell clone generation and expansion in both CRISPR-mediated knock-out (KO) and knock-in (KI) hiPSC lines. Using StemFlex medium and CloneR supplement in conjunction with Matrigel cell culture matrix, we show that cell viability and expansion during single-cell cloning in edited and unedited cells is significantly enhanced. Keeping all factors into account, we have successfully achieved hiPSC single-cell survival and cloning in both edited and unedited cells with rates as maximum as 70% in less than 2weeks. This simplified and efficient workflow will allow for a new level of sophistication in generating hiPSC-based disease models to promote rapid advancement in basic research and also the development of novel cellular therapeutics.

Efficient and iterative retron-mediated in vivo recombineering in Escherichia coli.

Recombineering is an important tool in gene editing, enabling fast, precise and highly specific in vivo modification of microbial genomes. Oligonucleotide-mediated recombineering via the in vivo production of single-stranded DNA can overcome the limitations of traditional recombineering methods that rely on the exogenous delivery of editing templates. By modifying a previously reported plasmid-based system for fully in vivo single-stranded DNA recombineering, we demonstrate iterative editing of independent loci by utilizing a temperature-sensitive origin of replication for easy curing of the editing plasmid from recombinant cells. Optimization of the promoters driving the expression of the system’s functional components, combined with targeted counterselection against unedited cells with Cas9 nuclease, enabled editing efficiencies of 90–100%. The addition of a dominant-negative mutL allele to the system allowed single-nucleotide edits that were otherwise unachievable due to mismatch repair. Finally, we tested alternative recombinases and found that efficiency significantly increased for some targets. Requiring only a single cloning step for retargeting, our system provides an easy-to-use method for rapid, efficient construction of desired mutants. Graphical

P104: BASE EDITING REPAIRS THE HBE MUTATION RESTORING THE PRODUCTION OF NORMAL GLOBIN CHAINS IN SEVERE HBE/β-THALASSEMIA PATIENT HEMATOPOIETIC STEM AND ERYTHROID CELLS

Aims: HbE/β-thalassemia is the commonest form of severe β-thalassemia, and comprises approximately 50% of all cases worldwide1. HbE/β-thalassemia is caused by the HbE codon 26 G>A mutation on one allele and any β0-thalassemia mutation on the other. There is a reduction in β-globin production, resulting in a relative excess in α-globin chains that leads to ineffective erythropoiesis. Importantly, individuals with a mutation on one, but not two, alleles have β-thalassemia trait, a carrier state with a normal phenotype shared by 1.5% of the world’s population2. Recent gene therapy and gene editing approaches have been developed to treat β-thalassemia but do not directly repair the causative mutation in-situ. Gene replacement approaches rely on lentiviral vector-based sequence insertion or homology directed repair (HDR). HbF induction strategies rely on non-homologous end joining targeting of enhancers in-trans. These approaches, whilst variably successful, are associated with potential safety concerns. Methods: Adenine base editors (ABEs) circumvent these problems by directly repairing pathogenic variants in-situ through deamination. ABEs catalyse A-T to G-C transitions. Conversion of the HbE codon to WT through base editing is an attractive strategy to recapitulate the phenotypically normal β-thalassemia trait state without potentially harmful double-strand breaks or random vector insertions. ABEs are able to convert the HbE codon (AAG) to wild-type (GAG), but also to GGG or AGG (Fig A). GGG at codon 26 is found in a naturally occurring haemoglobin, Hb Aubenas3. Heterozygotes have normal red cell indices and are phenotypically normal. We electroporated the latest generation of ABE8 editors4 as mRNA into 3 different severe HbE/β-thalassemia donor HSPCs with sgRNAs targeting the HbE codon. Results: The mean conversion from the HbE codon to a normal or normal variant in unselected cells was 86.2 (SD±8.1%, Fig B). The indel rate from inadvertent on-target Cas9 cleavage was below 0.5%. Edited cells did not show any perturbations in erythroid differentiation as assessed by Immunophenotyping and cellular morphology. In differentiated erythroid cells, RT-qPCR showed a mean fall in the α/β mRNA ratio to 0.65±0.08 (unedited patient cells normalised to 1, n=5, Figure C), indicating a reduction in the excess α-globin gene expression. Protein analysis by CE-HPLC showed a 3.6-fold reduction in HbE levels (SD±1.3) and a 13.5-fold increase in HbA/Hb Aubenas (SD±2.4, Fig D & E). In serial NSG-murine xenotransplantation experiments, base edited cells were found to persist in secondary transplants, showing editing is possible in long-term HSCs (mean editing efficiency 34.5%, Fig F). Potential off-target effects were assessed in-vitro by CIRCLE-seq5; most candidate sites were in intergenic and intronic regions (Fig G & H). The top 250 sites were sequenced using deep targeted NGS. Only 5 sites showed OT deaminations at low levels in patient cells (mean 1.5%). We developed a machine learning model to assess potential OT-effects on chromatin accessibility, at all candidate sites in 49 different blood cell types6. Only 17 potential edits were predicted to result in a significant change in chromatin state (Fig I). 3 of these were in the top 250 sites sequenced previously, and none showed deamination in-vivo. Conclusion: Together these data provide robust evidence for base editing being used as an effective and safe therapeutic strategy for HbE/β-thalassemia.

Holstein Friesian dairy cattle edited for diluted coat color as a potential adaptation to climate change

BackgroundHigh-producing Holstein Friesian dairy cattle have a characteristic black and white coat, often with large proportions of black. Compared to a light coat color, black absorbs more solar radiation which is a contributing factor to heat stress in cattle. To better adapt dairy cattle to rapidly warming climates, we aimed to lighten their coat color by genome editing.ResultsUsing gRNA/Cas9-mediated editing, we introduced a three bp deletion in the pre-melanosomal protein 17 gene (PMEL) proposed as causative variant for the semi-dominant color dilution phenotype observed in Galloway and Highland cattle. Calves generated from cells with homozygous edits revealed a strong color dilution effect. Instead of the characteristic black and white markings of control calves generated from unedited cells, the edited calves displayed a novel grey and white coat pattern.ConclusionThis, for the first time, verified the causative nature of the PMEL mutation for diluting the black coat color in cattle. Although only one of the calves was healthy at birth and later succumbed to a naval infection, the study showed the feasibility of generating such edited animals with the possibility to dissect the effects of the introgressed edit and other interfering allelic variants that might exist in individual cattle and accurately determine the impact of only the three bp change.