Inherited human Apollo deficiency causes severe bone marrow failure and developmental defects

Inherited bone marrow failure syndromes (IBMFS) represent a group of disorders typified by impaired production of one or several blood cell types. The telomere biology disorders dyskeratosis congenita (DC) and its severe variant Høyeraal-Hreidarsson (HH) syndrome are rare IBMFS characterized by bone marrow failure, developmental defects, and various premature aging complications associated with critically short telomeres. Here we identified biallelic variants in the gene encoding the 5'-to-3' DNA exonuclease Apollo/SNM1B in three unrelated patients presenting with a DC/HH phenotype consisting of early onset hypocellular bone marrow failure, B and NK lymphopenia, developmental anomalies, microcephaly and/or intrauterine growth retardation. All three patients carry a homozygous or compound heterozygous (in combination with a null-allele) missense variant affecting the same residue L142 (L142F or L142S) located in the catalytic domain of Apollo. Apollo-deficient cells from patients exhibited spontaneous chromosome instability and impaired DNA repair that was complemented by CRISPR/Cas9-mediated gene correction. Furthermore, patients' cells showed signs of telomere fragility that were however not associated with global reduction of telomere length. Unlike patients' cells, human Apollo KO HT1080-cell lines showed strong telomere dysfunction accompanied by excessive telomere shortening, suggesting that the L142S and L142F Apollo variants are hypomorphic. Collectively, these findings define human Apollo as a genome caretaker and identify biallelic Apollo variants as a genetic cause of a hitherto unrecognized severe IBMFS combining clinical hallmarks of DC/HH with normal telomere length.

Impact of Formulation Conditions on Lipid Nanoparticle Characteristics and Functional Delivery of CRISPR RNP for Gene Knock-Out and Correction

The CRISPR-Cas9 system is an emerging therapeutic tool with the potential to correct diverse ge-netic disorders. However, for gene therapy applications an efficient delivery vehicle is required, capable of delivering the CRISPR-Cas9 components into the cytosol of the intended target cell population. Once there, the ribonucleoprotein complex (RNP) can be transported into the nucleus. Lipid nanoparticles (LNP) serve as promising candidates for delivery of CRISPR-Cas9 RNP. These delivery vehicles have been optimized for the delivery of nucleic acids, such as mRNA. Co-delivery of Cas9 encoding mRNA with the accompanying sgRNA leads to translation of the Cas9 protein and formation of the Cas9 RNP inside the cell. Only recently, direct delivery of the CRISPR-Cas9 RNP complexes has been explored, which requires adjustments to the LNP formulation. In this study, the importance of buffer composition and cationic charge during RNP and ssDNA en-trapment in LNP are demonstrated. After optimizing several formulation parameters, LNP were prepared that were colloidally stable in human plasma and efficiently deliver the SpCas9 RNP and ssDNA for HDR-correction in reporter cells. Under optimal formulation conditions, gene knock-out and gene correction efficiencies as high as 80% and 20%, respectively were achieved at nanomolar CRISPR-Cas9 RNP concentrations.

A selectable all-in-one CRISPR prime editing piggyBac transposon allows for highly efficient gene editing in human cell lines

CRISPR prime-editors are emergent tools for genome editing and offer a versatile alternative approach to HDR-based genome engineering or DNA base-editors. However, sufficient prime-editor expression levels and availability of optimized transfection protocols may affect editing efficiencies, especially in hard-to-transfect cells like hiPSC. Here, we show that piggyBac prime-editing (PB-PE) allows for sustained expression of prime-editors. We demonstrate proof-of-concept for PB-PE in a newly designed lentiviral traffic light reporter, which allows for estimation of gene correction and defective editing resulting in indels, based on expression of two different fluorophores. PB-PE can prime-edit more than 50% of hiPSC cells after antibiotic selection. We also show that improper design of pegRNA cannot simply be overcome by extended expression, but PB-PE allows for estimation of effectiveness of selected pegRNAs after few days of cultivation time. Finally, we implemented PB-PE for efficient editing of an amyotrophic lateral sclerosis-associated mutation in the SOD1-gene of patient-derived hiPSC. Progress of genome editing can be monitored by Sanger-sequencing, whereas PB-PE vectors can be removed after editing and excised cells can be enriched by fialuridine selection. Together, we present an efficient prime-editing toolbox, which can be robustly used in a variety of cell lines even when non-optimized transfection-protocols are applied.

S. Aureus Cas9 Leads to Higher Sickle Mutation Correction Compared to S. Pyogenes Cas9 in Patient Hematopoietic Stem and Progenitor Cells

Introduction: Sickle cell disease (SCD) is a red blood cell disorder caused by a single nucleotide mutation in the β-globin gene (HBB). Allogeneic hematopoietic stem cell transplantation (HSCT) is the only available cure, but is available to only a minority of patients and can be associated with high morbidity and mortality. CRISPR/Cas9 mediated genome editing may provide a permanent cure for SCD patients by correcting the sickle mutation in HBB in hematopoietic stem and progenitor cells (HSPCs). Previously, we achieved ~39% sickle mutation correction in SCD HSPCs by delivering S. pyogenes (Spy) Cas9/R-66S gRNA as ribonucleoprotein (RNP) and single-stranded oligodeoxynucleotides (ssODN) corrective donor template. S. aureus (Sau) Cas9 has potentially advantageous properties to improve therapeutic gene editing efficiency and safety, including smaller size allowing for efficient in vivo delivery and longer Protospacer Adjacent Motif (PAM) sequence for higher specificity. However, although in general, the cutting efficiency of SauCas9 is lower than SpyCas9, the differences in gene correction and other gene-editing outcomes between SpyCas9 and SauCas9 have not been well studied. Methods: With our R-66S gRNA sequence targeting the sickle mutation, the PAM sequence of SauCas9 (NGGRRT) is mutually permissive with that of SpyCas9 (NGG), allowing the same sequence to be targeted by both Cas9 nucleases. We delivered R-66S gRNA with SpyCas9 and SauCas9 respectively as RNP, along with corrective ssODN donor template into SCD HSPCs. We analyzed sickle mutation correction rate and small insertions and deletions (INDELs) profile by Next Generation Sequencing (NGS). Results/discussions: We found that although the INDEL rate of SpyCas9 is higher than SauCas9 at the same molar concentration of RNP, SauCas9 gave 43% sickle mutation correction, slightly higher than SpyCas9 (39%), demonstrating efficient homology-directed repair (HDR) mediated gene correction by SauCas9. To further investigate the potential for clinical translation, we will perform in-depth efficiency and safety characterization comparing SauCas9 and SpyCas9 mediated sickle mutation correction therapy in SCD HSPCs. Conclusion: In this work, we showed that, compared with the highly-optimized and widely-used SpyCas9, SauCas9 leads to a higher sickle mutation correction in SCD HSPCs, demonstrating the therapeutic potential of SauCas9 for treating SCD. We will further investigate the efficiency and safety of gene-edited therapy mediated by these two Cas9 orthologs, including in-depth characterization of off-target effects, chromosomal rearrangement and aberrations, and large genomic modifications. We will differentiate gene-corrected SCD HSPCs to study erythropoiesis and red cell phenotype, including normal hemoglobin production and reduced sickling under hypoxic conditions. Lastly, we will evaluate the engraftment efficiency of gene-edited cells in Nonirradiated NOD,B6.SCID Il2rγ -/- Kit (W41/W41) (NBSGW) mice that support the engraftment of human hematopoietic stem cells. Disclosures Sheehan: Forma Therapeutics: Research Funding; Beam Therapeutics: Research Funding; Novartis: Research Funding.

Cedar Trial in Progress: A First in Human, Phase 1/2 Study of the Correction of a Single Nucleotide Mutation in Autologous HSCs (GPH101) to Convert HbS to HbA for Treating Severe SCD

Background Sickle cell disease (SCD) is a recessive monogenic disease caused by a single point mutation in which glutamic acid replaces valine in Codon 6 of the human beta-globin gene (HBB) leading to the production of abnormal globin chains (HbS) that polymerize and cause erythrocytes to sickle. This results in hemolytic anemia, vaso-occlusion and organ damage, which leads to lifelong complications and early mortality. Allogeneic hematopoietic stem cell transplant (allo-HSCT) is the only known cure for SCD, however, its use is limited by the lack of well-matched donors, need for immunosuppression, risk of graft versus host disease and graft rejection. GPH101 is an investigational, autologous, hematopoietic stem cell (HSC) drug product (DP) designed to correct the SCD mutation in the HBB gene ex vivo using a high fidelity Cas9 (CRISPR associated protein 9) paired with an AAV6 (adeno-associated virus type 6) delivery template, efficiently harnessing the natural homology directed repair (HDR) cellular pathway. This approach has the potential to restore normal adult hemoglobin (HbA) production while simultaneously reducing HbS levels. In preclinical studies, HBB gene correction in SCD donor HSCs resulted in ≥60% of gene-corrected alleles in vitro with minimal off-target effects. Gene corrected cells were successfully differentiated toward the erythroid lineage and produced ≥70% HbA in vitro. Long-term engraftment of gene-corrected HSCs was demonstrated in vivo, following transplant into immunodeficient mice, with multi-lineage allelic gene correction frequencies well above the predicted curative threshold of 20%, with potential of this approach to be equivalent or superior to allo-HSCT. In addition, HSC-based correction in an SCD mouse model led to stable adult hemoglobin production, increased erythrocyte lifespan and reduction in sickling morphology, demonstrating the therapeutic potential of this gene correction platform as a curative approach in SCD. Study Design and Methods CEDAR (NCT04819841) is a first-in-human, open-label, single-dose, multi-site Phase 1/2 clinical trial in participants with severe SCD designed to evaluate safety, efficacy and pharmacodynamics (PD) of GPH101. Approximately 15 adult (18-40 years) and adolescent (12-17 years) participants will be enrolled across 5 sites, with adolescent enrollment proceeding after a favorable assessment of adult safety data by a Safety Monitoring Committee. Participants must have a diagnosis of severe SCD (βS/βS), defined as ≥ 4 severe vaso-occlusive crises (VOCs) in the 2 years prior and/or ≥ 2 episodes of acute chest syndrome (ACS), in 2 years prior with at least 1 episode in the past year. Participants on chronic transfusion therapy may be eligible if required VOC and ACS criteria are met in the 2 years prior to the initiation of transfusions. Key exclusion criteria include availability of a 10/10 human leukocyte antigen-matched sibling donor, or prior receipt of HSCT or gene therapy. After eligibility confirmation including screening for pre-treatment cytogenetic abnormalities, participants will undergo plerixafor mobilization and apheresis, followed by CD34+ cell enrichment and cryopreservation, undertaken locally at each trial site before shipment to a centralized manufacturer for GPH101 production. After GPH101 release, participants will undergo eligibility reconfirmation prior to busulfan conditioning and DP infusion. Safety, efficacy and PD measurements will occur for 2 years post-infusion; a long-term follow up study will be offered to participants for an additional 13 years of monitoring. The primary endpoint for this study is safety, measured by the kinetics of HSC engraftment, transplant related mortality, overall survival and frequency and severity of adverse events. Secondary endpoints will explore efficacy and PD, including levels of globin expression as compared to baseline, gene correction rates, clinical manifestations of SCD (including VOC and ACS), laboratory parameters, complications and organ function. In addition, cerebral hemodynamics and oxygen delivery will be assessed by magnetic resonance techniques. Key exploratory endpoints include evaluation of patient-reported outcomes, erythrocyte function, on-target and off-target editing rates, and change from baseline in select SCD characteristics. Disclosures Kanter: Fulcrum Therapeutics, Inc.: Consultancy; Novartis: Consultancy, Honoraria, Membership on an entity's Board of Directors or advisory committees; Forma: Consultancy, Honoraria, Membership on an entity's Board of Directors or advisory committees; Agios: Honoraria, Membership on an entity's Board of Directors or advisory committees; Beam: Honoraria, Membership on an entity's Board of Directors or advisory committees; Sanofi: Honoraria, Membership on an entity's Board of Directors or advisory committees; Graphite Bio: Consultancy; GuidePoint Global: Honoraria; Fulcrum Tx: Consultancy. Thompson: Agios Pharmaceuticals: Consultancy; Graphite Bio: Research Funding; Vertex: Research Funding; Beam Therapeutics: Consultancy; Celgene: Consultancy, Research Funding; Biomarin: Research Funding; Baxalta: Research Funding; CRISPR Therapeutics: Research Funding; Global Blood Therapeutics: Current equity holder in publicly-traded company; bluebird bio: Consultancy, Research Funding; Novartis: Research Funding. Porteus: Versant Ventures: Consultancy; CRISPR Therapeutics: Current equity holder in publicly-traded company; Allogene Therapeutics: Current equity holder in publicly-traded company, Membership on an entity's Board of Directors or advisory committees; Ziopharm: Current equity holder in publicly-traded company, Membership on an entity's Board of Directors or advisory committees; Graphite Bio: Current equity holder in publicly-traded company, Membership on an entity's Board of Directors or advisory committees. Intondi: Graphite Bio: Current Employment, Current equity holder in publicly-traded company; Global Blood Therapeutics: Current equity holder in publicly-traded company, Ended employment in the past 24 months. Lahiri: Graphite Bio: Current Employment, Current equity holder in publicly-traded company. Dever: Graphite Bio: Current Employment, Current equity holder in publicly-traded company. Petrusich: bluebird bio: Current equity holder in publicly-traded company, Ended employment in the past 24 months; Graphite Bio: Current Employment, Current equity holder in publicly-traded company. Lehrer-Graiwer: Global Blood Therapeutics: Current equity holder in publicly-traded company, Ended employment in the past 24 months; Graphite Bio: Current Employment, Current equity holder in publicly-traded company.

Engineered PAM-flexible FnCas9 variants for robust and specific genome editing and diagnostics

The clinical success of CRISPR therapies is dependent on the safety and efficacy ofCas proteins. The Cas9 from Francisella novicida (FnCas9) has negligible affinity formismatched substrates enabling it to discriminate off-targets in DNA with very highprecision even at the level of binding. However, its cellular targeting efficiency is low,limiting its use in therapeutic applications. Here, we rationally engineer the protein todevelop engineered(enFnCas9) variants with enhanced activity and expand its cellularediting activity to genomic loci previously inaccessible. Notably, some of the variantsrelease the protospacer adjacent motif (PAM) constraint from NGG to NGR/NRGmaking them rank just below SpCas9-RY and SpCas9-NG in their accessibility acrosshuman genomic sites. The enFnCas9 proteins, similar to Cas12a and Cas12f, harborhigh intrinsic specificity and can diagnose single nucleotide variants accurately.Importantly, they provide superior outcomes in terms of editing efficiency, knock-inrates, and off-target specificity over other engineered high-fidelity versions of SpCas9(SpCas9-HF1 and eSpCas9). Broad targeting range coupled with remarkablespecificity of DNA interrogation underscores the utility of these variants for safe andefficient therapeutic gene correction across multiple cell lines and target loci.

Rhesus Macaques As a Preclinical Model Organism for RUNX1-FPD

Germline loss-of-function (LOF) heterozygous mutations in the central hematopoietic transcription factor RUNX1 gene cause the marrow failure/malignancy predisposition syndrome Familial Platelet Disorder with associated Myeloid Malignancies (FPDMM, or FPD). Patients with FPD have defective megakaryocytic development, low platelet counts, and a very high (35-50%) life-long risk of hematological malignancies, particularly MDS/AML. Murine heterozygous gene knockout models do not recapitulate the human phenotype in terms of thrombocytopenia or myeloid leukemia progression. Although gene correction of the RUNX1 mutation in hematopoietic stem and progenitor cells (HSPCs) is being considered as a possible treatment approach, it is unknown whether mutation-corrected HSPCs will have the hoped for advantage over RUNX1 mutant HSCs in vivo, likely necessary to significantly lower leukemia risk. In order to study the relative function of wildtype and RUNX1-mutated HSPCs in vivo in a model with close hematopoietic similarity to humans, we generated a rhesus macaque FPD competitive repopulation model via CRISPR/Cas9 NHEJ editing of the RUNX1 gene versus the AAVS1 safe-harbor control locus. We transplanted mixtures of autologous HSPCs edited at the two loci: 75% RUNX1-edited/25% AAVS1- edited CD34+ HSPCs in animal 1 and 25% RUNX1-edited/75% AAVS1-edited CD34+ HSPCs in animal 2, following conditioning with total body irradiation. Both animals engrafted tri-lineage hematopoiesis promptly following transplantation. However, platelet numbers remained below the normal range long-term in animal 1 receiving a higher ration of RUNX1-edited HSPCS and below counts of macaques receiving HSPCs edited at other loci (Figure 1). Bone marrow morphology at 6 months was normal. To assess the HSPC function of RUNX1 mutant versus AAV1 control and unedited WT cells we tracked RUNX1 and AAVS1-mutated allele frequencies in blood cells over time via deep sequencing (Figure 2). In the infusion products (IP), allele fractions reflected the desired ratios. In both animals, AAVS1-edited cells dominated compared to RUNX1-edited cells. However, in animal 1, RUNX1-mutated cells expanded over time eventually exceeding the ratio in the IP, and in animal 2, levels of RUNX1 and AAVS1-mutated cells were equivalent long-term. Marrow analyzed at 6 months showed heterozygous RUNX1-mutated CFU at levels concordant with mutation frequencies in the blood, but no homozygous RUNX1 mutated CFU, suggesting homozygous LOF is not compatible with long-term HSPC function. In conclusion, we have created pre-clinical model for FPD via CRISPR/Cas editing of HSPCs in rhesus macaques. The lack of a competitive advantage for wildtype or control-locus edited HSPCs over RUNX1 heterozygous-mutated HSPCs long-term in our model suggests that gene correction approaches for FPD will be challenging, particularly to reverse the MDS/AML predisposition phenotype. Figure 1 Figure 1. Disclosures No relevant conflicts of interest to declare.

Directed Differentiation of Human Pluripotent Stem Cells toward Skeletal Myogenic Progenitors and Their Purification Using Surface Markers

Advancements in reprogramming somatic cells into induced pluripotent stem cells (iPSCs) have provided a strong framework for in vitro disease modeling, gene correction and stem cell-based regenerative medicine. In cases of skeletal muscle disorders, iPSCs can be used for the generation of skeletal muscle progenitors to study disease mechanisms, or implementation for the treatment of muscle disorders. We have recently developed an improved directed differentiation method for the derivation of skeletal myogenic progenitors from hiPSCs. This method allows for a short-term (2 weeks) and efficient skeletal myogenic induction (45–65% of the cells) in human pluripotent stem cells (ESCs/iPSCs) using small molecules to induce mesoderm and subsequently myotomal progenitors, without the need for any gene integration or modification. After initial differentiation, skeletal myogenic progenitors can be purified from unwanted cells using surface markers (CD10+CD24−). These myogenic progenitors have been extensively characterized using in vitro gene expression/differentiation profiling as well as in vivo engraftment studies in dystrophic (mdx) and muscle injury (VML) rodent models and have been proven to be able to engraft and form mature myofibers as well as seeding muscle stem cells. The current protocol describes a detailed, step-by-step guide for this method and outlines important experimental details and troubleshooting points for its application in any human pluripotent stem cells.

CRISPR/Cas9 Delivery System Engineering for Genome Editing in Therapeutic Applications

The clustered regularly interspaced short palindromic repeats (CRISPR)/associated protein 9 (CRISPR/Cas9) systems have emerged as a robust and versatile genome editing platform for gene correction, transcriptional regulation, disease modeling, and nucleic acids imaging. However, the insufficient transfection and off-target risks have seriously hampered the potential biomedical applications of CRISPR/Cas9 technology. Herein, we review the recent progress towards CRISPR/Cas9 system delivery based on viral and non-viral vectors. We summarize the CRISPR/Cas9-inspired clinical trials and analyze the CRISPR/Cas9 delivery technology applied in the trials. The rational-designed non-viral vectors for delivering three typical forms of CRISPR/Cas9 system, including plasmid DNA (pDNA), mRNA, and ribonucleoprotein (RNP, Cas9 protein complexed with gRNA) were highlighted in this review. The vector-derived strategies to tackle the off-target concerns were further discussed. Moreover, we consider the challenges and prospects to realize the clinical potential of CRISPR/Cas9-based genome editing.