ABSTRACT:Sickle cell disease (SCD) is a red blood cell disorder caused by a mutation in the β-globin gene, leading to sickle hemoglobin polymerization under low oxygen conditions. Both CRISPR-Cas9 editing and lentiviral transduction have shown promising clinical outcomes, but it remains unclear which approach is superior. Alternatively, new editing tools such as base editing may also be promising and reduce risks of genotoxicity. To compare these approaches, we studied them in an immunocompromised mouse model. We optimized ex vivo conditions in CD34+ hematopoietic stem and progenitor cells (HSPC) and infused edited SCD HSPC into busulfan-conditioned nonirradiated NOD,B6.SCID Il2rγ-/-KitW41/W41 (NBSGW) mice. Ex vivo analysis confirmed successful editing and transduction. At 16 weeks, bone marrow analysis showed similar human CD45+ cell engraftment across all groups (75%-90%). In the competitive transplantation group, there was a lower amount of B-cell lymphoma/leukemia 11A enhancer editing than base editing and lentiviral transduction. A secondary transplantation model yielded similar results. An antisickling assay showed significantly higher red blood cell sickling reduction in the base editing, transduction, and competitive transplantation groups compared to CRISPR-Cas9. In conclusion, although all methods showed therapeutic potential, base editing and lentiviral transduction provided superior outcomes over CRISPR-Cas9-mediated editing in a competitive murine transplantation model.

Hematopoietic stem cells (HSCs) reconstitute blood cells throughout life. DNA-level correction of HSCs allows for a one-time cure of genetic diseases, including sickle cell disease (SCD). Sickle cell disease is one of the most common single-gene disorders; therefore, SCD is a prime candidate for gene therapy. Several drug therapies are available for SCD, including hydroxyurea, which is the first-line choice despite requiring lifelong administration. Allogeneic HSC transplantation is a one-time, curative treatment for SCD with limited availability of histocompatible donors. Therefore, autologous HSC gene therapy was developed using patients' own HSCs with lentiviral gene addition/silencing and clustered regularly interspaced short palindromic repeats gene editing, making gene therapy applicable to most patients. However, the established method of HSC gene therapy requires costly and complex ex vivo HSC culture. Therefore, in vivo HSC gene therapy is being developed to treat SCD, envisioning a single-injection HSC-targeted gene delivery system. This review discusses various therapeutic methods to treat SCD, the development of HSC gene therapy, and clinical gene therapy trials in SCD, ranging from FDA-approved to novel in vivo gene therapy.

Sickle cell disease (SCD) is a single-gene disorder caused by a point mutation of the β-globin gene, resulting in hemolytic anemia, acute pain, multiorgan damage, and early mortality. Hydroxyurea is a first-line drug therapy that switches sickle-globin to non-pathogenic γ-globin; however, it requires lifelong oral administration. Allogeneic hematopoietic stem cell (HSC) transplantation allows for a one-time cure for SCD, albeit with histocompatibility limitations. Therefore, autologous HSC gene therapy was developed to cure SCD in a single treatment, without HSC donors. Current HSC gene therapy is based on the ex vivo culture of patients’ HSCs with lentiviral gene addition and gene editing, followed by autologous transplantation back to the patient. However, the complexity of the treatment process and high costs hinder the universal application of ex vivo gene therapy. Therefore, the development of in vivo HSC gene therapy, where gene therapy tools are directly administered to patients, is desirable to provide a more accessible, cost-effective solution that can cure SCD worldwide. In this review, we discuss current treatments, including drug therapies, HSC transplantation, and ex vivo gene therapy; the development of gene therapy tools; and progress toward curative in vivo gene therapy in SCD.

Editing the +58 region of the BCL11A erythroid enhancer has shown promise in treating β-globin disorders. To address variations in fetal hemoglobin (HbF) response, we investigated editing both +58 and +55 enhancers. Rhesus macaques transplanted with edited hematopoietic stem/progenitor cells (HSPCs) following busulfan conditioning exhibited durable, high-level (∼90%) editing frequencies post transplantation with sustained HbF reactivation over 4 years, without hematological perturbations. HbF levels were further boosted by stress erythropoiesis or hydroxyurea. Bone marrow analysis revealed that gene edits were predominantly programmed deletions, programmed inversions, and short indels, each disrupting the enhancer core TGN7-9WGATAR half E-box/GATA binding motifs. Nonprogrammed long deletions were disfavored in engrafting cells. CD45 antibody-drug conjugate (ADC) conditioning achieved comparable engraftment and HbF reactivation, whereas lentiviral vector tracking showed polyclonal reconstitution with dynamics similar to animals conditioned with total body irradiation (TBI) or busulfan. Joining CD45-ADC conditioning with combined enhancer editing presents an effective strategy for β-hemoglobinopathies, enabling durable HbF reactivation without chemotherapy.

Sickle cell disease (SCD) is a common, severe genetic blood disorder. Current pharmacotherapies are partially effective and allogeneic hematopoietic stem cell transplantation is associated with immune toxicities. Genome editing of patient hematopoietic stem cells (HSCs) to reactivate fetal hemoglobin (HbF) in erythroid progeny offers an alternative potentially curative approach to treat SCD. Although the FDA released guidelines for evaluating genome editing risks, it remains unclear how best to approach pre-clinical assessment of genome-edited cell products. Here, we describe rigorous pre-clinical development of a therapeutic γ-globin gene promoter editing strategy that supported an investigational new drug application cleared by the FDA. We compared γ-globin promoter and BCL11A enhancer targets, identified a potent HbF-inducing lead candidate, and tested our approach in mobilized CD34+ hematopoietic stem progenitor cells (HSPCs) from SCD patients. We observed efficient editing, HbF induction to predicted therapeutic levels, and reduced sickling. With single-cell analyses, we defined the heterogeneity of HbF induction and HBG1/HBG2 transcription. With CHANGE-seq for sensitive and unbiased off-target discovery followed by targeted sequencing, we did not detect off-target activity in edited HSPCs. Our study provides a blueprint for translating new ex vivo HSC genome editing strategies toward clinical trials for treating SCD and other blood disorders.

BIVV003 is a gene-edited autologous cell therapy in clinical development for the potential treatment of sickle cell disease (SCD). Hematopoietic stem cells (HSC) are genetically modified with mRNA encoding zinc finger nucleases (ZFN) that target and disrupt a specific regulatory GATAA motif in the BCL11A erythroid enhancer to reactivate fetal hemoglobin (HbF). We characterized ZFN-edited HSC from healthy donors and donors with SCD. Results of preclinical studies show that ZFN-mediated editing is highly efficient, with enriched biallelic editing and high frequency of on-target indels, producing HSC capable of long-term multilineage engraftment in vivo, and express HbF in erythroid progeny. Interim results from the Phase 1/2 PRECIZN-1 study demonstrated that BIVV003 was well-tolerated in seven participants with SCD, of whom five of the six with more than 3 months of follow-up displayed increased total hemoglobin and HbF, and no severe vaso-occlusive crises. Our data suggest BIVV003 represents a compelling and novel cell therapy for the potential treatment of SCD.

Ex vivo resting culture is a standard procedure following genome editing in hematopoietic stem and progenitor cells (HSPCs). However, prolonged culture may critically affect cell viability and stem cell function. We investigated whether varying durations of culture resting times impact the engraftment efficiency of human CD34+ HSPCs edited at the BCL11A enhancer, a key regulator in the expression of fetal hemoglobin. We employed electroporation to introduce CRISPR-Cas9 components for BCL11A enhancer editing and compared outcomes with non-electroporated (NEP) and electroporated-only (EP) control groups. Post-electroporation, we monitored cell viability, death rates, and the frequency of enriched hematopoietic stem cell (HSC) fractions (CD34+CD90+CD45RA- cells) over a 48-hour period. Our findings reveal that while the NEP group showed an increase in cell numbers 24 hours post-electroporation, both EP and BCL11A-edited groups experienced significant cell loss. Although CD34+ cell frequency remained high in all groups for up to 48 hours post-electroporation, the frequency of the HSC-enriched fraction was significantly lower in the EP and edited groups compared to the NEP group. In NBSGW xenograft mouse models, both conditioned with busulfan and non-conditioned, we found that immediate transplantation post-electroporation led to enhanced engraftment without compromising editing efficiency. Human glycophorin A+ (GPA+) red blood cells (RBCs) sorted from bone marrow of all BCL11A edited mice exhibited similar levels of γ-globin expression, regardless of infusion time. Our findings underscore the critical importance of optimizing the culture duration between genome editing and transplantation. Minimizing this interval may significantly enhance engraftment success and minimize cell loss without compromising editing efficiency. These insights offer a pathway to improve the success rates of genome editing in HSPCs, particularly for conditions like sickle cell disease.

Gene therapy targeting hematopoietic stem cells (HSCs) is a promising treatment for a variety of genetic disorders, including immunodeficiency, hemoglobinopathies, congenital cytopenia, and metabolic diseases. HSCs can reconstitute peripheral blood throughout life due to their capacity for self-renewal and their hematopoietic multipotency. This makes it possible to cure genetic diseases for an entire lifetime by replacing or repairing pathogenic mutations/deletions in HSCs. Autologous HSC-targeted gene therapies entailing lentiviral gene addition as well as gene editing are currently under development. These can be widely applied to most patients, as there is no requirement for a suitable donor. Current gene addition/editing therapies are based on harvesting the patient's CD34+ HSCs, performing gene modification ex vivo, and then transplanting the modified HSCs back into the patient. The efficacy of ex vivo lentiviral HSC gene therapy has been proved in recent trials; however, the ex vivo process requires a GMP-level cell processing center and is expensive, which limits its global application. It is therefore crucial to develop in vivo HSC gene therapies, in which a therapeutic gene or gene editing tools can be delivered directly into bone marrow HSCs via systemic administration without ex vivo culture. This manuscript presents an overview of the current HSC-targeted gene therapies using lentiviral vectors.

Hematopoietic stem cell (HSC)-targeted gene therapy is curative for various genetic blood diseases, and its efficacy has been demonstrated in recent clinical trials. HSCs have self-renewal and hematopoietic multipotency; therefore, repairing pathological mutations or defects in HSCs allows for a lifelong cure with a single treatment. Autologous HSC gene therapy has been developed by lentiviral gene addition or gene editing, and is an option for most patients because it does not require a compatible donor. Current HSC gene therapy is based on ex vivo methods, in which patient HSCs are harvested, genetically modified ex vivo, and autologously transplanted into patients. However, the complexity of this process and the high cost of treatment are hindering the spread of gene therapy. Therefore, in vivo HSC gene therapy is being developed to deliver gene therapy tools directly into bone marrow HSCs by administration without ex vivo culture.

Current genetic therapies for sickle cell disease (SCD) require busulfan or other myeloablative conditioning regimens prior to cell infusion. Busulfan leads to both acute and chronic toxicities including infertility, organ injury and the potential for increased rates of malignancy. Due to the significant adverse effects of myeloablative conditioning with busulfan or other conditioning agents, many patients will not be eligible or will not choose to receive a potentially transformative therapy. To address this challenge, we developed a non-genotoxic conditioning strategy with our Engineered Stem Cell Antibody Paired Evasion (ESCAPE) approach where we epitope engineered the hematopoietic stem and progenitor cell (HSPC) surface protein CD117 (cKIT) with a base editor to retain normal CD117 receptor function, but escape recognition by a cognate monoclonal antibody (mAb) that recognizes the wildtype CD117 protein and can be used for conditioning and in vivo selection of edited cells. The ESCAPE CD117 edit is combined with a therapeutic target edit in the γ-globin gene (HBG1/2) promoters to induce high levels of fetal hemoglobin (HbF) to prevent polymerization of sickle hemoglobin and the associated pathophysiology of SCD. This ESCAPE approach enables elimination of the toxic effects of myeloablative conditioning and provides a selective advantage for edited cells as the mAb does not need to be cleared before or following infusion of edited cells. We demonstrated the ESCAPE concept in humanized wild type (WT) NBSGW mice treated with CD117 mAb followed by transplantation with multiplex edited human HSPCs. This conditioning/transplant strategy resulted in >50% HbF in sorted bone marrow GlyA positive cells, indicating the potential for significant therapeutic benefit. Highlighting the exquisite dependency of erythropoiesis on CD117 signaling, we also observed that ex vivo CD117 mAb treatment of multiplex edited HSPCs led to rapid selection of multiplex edited erythroid cells in mixtures of unedited and edited cells. To investigate whether ESCAPE, incorporating single agent CD117 mAb conditioning, could achieve rapid and robust induction of HbF in an immunocompetent host, we used a rhesus macaque (Macaca mulatta) autologous CD34+ HSCT model. First, we demonstrated that the CD117 mAb was cross-reactive and bound rhesus HSPCs to induce apoptosis ex vivo. Mobilized rhesus CD34+ HSPCs were then multiplex edited for CD117 and HBG1/2 and HbF levels of >50% were confirmed in the cell product. Successful engraftment was achieved in a busulfan-conditioned animal, resulting in peripheral blood F-cell levels up to 98% and γ-globin levels up to 88%. We next employed the ESCAPE multiplex-edited autologous CD34+ HSPC approach in two additional rhesus macaques conditioned with 10 mg/kg or 25 mg/kg of CD117 mAb prior to HSCT. Subsequently, these animals received additional mAb treatments post-transplantation to provide a competitive advantage for edited cells with WT CD117 receptor occupancy maintained at 80-90%. In contrast to the busulfan treated animal, the mAb was well tolerated at both doses and no supportive care was necessary for the mAb conditioned animals. We observed significant induction of HbF at early timepoints post-transplant. F-cell levels rose to 61% in the periphery as early as 8 weeks post-transplant in one of the non-human primates (NHPs), and both NHPs stabilized at ~85% F-cells at week 35. Concomitant early induction of γ-globin was observed in these NHPs with ~37% γ-globin levels achieved at 8 weeks post-transplant and reaching ~55% at 35 weeks. Two nontransplant control NHPs received 10 mg/kg mAb dosing and did not demonstrate HbF induction (< 1% total γ-globin). Our findings suggest that the ESCAPE approach, enabled by multiplex base editing and using CD117 mAb conditioning and selection, effectively reactivates HbF production without the need for genotoxic conditioning or post-transplant supportive measures and hematological perturbations in NHPs. This work lays the groundwork for potential therapeutic advancements in SCD treatment that avoid busulfan or other myeloablative conditioning related morbidities, enable rapid induction of therapeutically relevant levels of HbF and brings us closer to the goal of advancing gene therapy to SCD patients across the full spectrum of disease severity, including in resource limited environments.

Sickle cell disease (SCD) is a monogenic hemoglobin (Hgb) disorder characterized by a single pointmutation in the β-globin gene which causes Hgb polymerization under low oxygen tension and sickling of red blood cells (RBCs) which in turn drives severe systemic injury. Gene addition technologies like hematopoietic stem cell (HSC) targeted lentiviral (LV) gene therapy are among the most promising curative strategies for SCD. To expand this approach, improvements in LV titer, transduction efficiency, along with suppression of native sickle-globin, are preferred to generate successful therapeutic outcomes. One such improvement targets LV orientation: the relationship between the DNA encoding the β-globin gene in the LV and the promoter driving healthy β-globin gene expression. We created a forward-oriented β-globin-expression vector with an improved titer that results in more efficient transduction of human CD34+ cells in vitro, in human reconstituting CD34+ cells in xenografted mice, and in rhesus reconstituting CD34+ cells(Uchida N. et al., 2019). We report here on further optimization of LVs that in addition to forward orientation, also express shRNA that decreases native β-globin and sickle-globin, which could enhance the assembly of corrected Hgb tetramers. Seven unique constructs which varied in anti-β-globin shRNA sequences, packaging elements, and regulatory elements, were transduced into MEL-BB88 cells and expanded for 2 weeks. gDNA was collected for vector copy number (VCN) analysis, and treated with 10 µM hemin to induce Hgb expression. Five days after hemin-induction, cells were harvested for benzidine staining and immunoblot analysis to validate globin expression. Sca-1+ HSC/progenitor (HSPC) BM cells were isolated from Towne's humanized SCD donor mice prior to stimulation for 48 h with cytokines, then transduced at an MOI of 24 twice, 24 h apart. To assess the in vivo engraftment potential of the transduced HSPCs, 2.5x10^6 cells were transplanted into lethally irradiated CD45.1 recipient mice. Peripheral blood (PB) was sampled at 2-week intervals for PBMC chimerism and lineage engraftment, Hgb electrophoresis, Lorrca OxygenScan and OsmoScan assays, and PB VCN. Whole BM from engrafted recipients was collected at 30 weeks and used for secondary transplantation assays which were again analyzed over 24 weeks for PB engraftment, VCN, and donor Hgb expression. LVs were produced at varying titers and all successfully transduced MEL-BB88 cells with VCNs between 1 and 5 that were maintained following hemin induction. Interestingly, variations in RRE elements dramatically affected human globin expression in vitro. Furthermore, the transplanted mice engrafted fully with HSPCs transduced with multiple vectors. Notably, mice transplanted with HSPCs transduced with vector CX0066p (LCR-Enhancer-P-HBB-shmiR-3g1.223 + 3g2.30a-R1) maintained a VCN of 0.2-0.4 in PB for the 30-week duration of the primary transplant. Cellulose acetate gels of RBCs from PB showed ~40% HbA in a background of HbS. The P50 right-shifted towards normal, RBC counts increased, WBC counts decreased, and OxygenScan parameters were all corrected towards controls and away from those values obtained from mice transplanted with mock transduced HSPCs. EI max and EI min increased while pO2@95% decreased. Lastly, the RBC sickling was corrected in an in vitro sickling assay. In conclusion, based on LV titer, MEL cell expression, transduction efficiency, engraftment of modified cells, in vivo globin expression, anti-sickling activity, and oxygen transport, these constructs provide outstanding Hgb expression and phenotype correction. Consequently, these candidates serve as a solid foundation for future human gene therapy products in the treatment of SCD.