BackgroundOn-demand gene editing of autologous cells offers a novel therapeutic option for inborn errors of immunity (IEIs) caused by dominant-negative variants, which are often poorly addressed by conventional viral gene addition or hematopoietic stem cell transplantation (HSCT). IRF4 encodes a transcription factor critical for T cell differentiation, Th17 polarization, memory formation, and tissue homing via integrins such as α4β7. Heterozygous neomorphic variants at T95 disrupt IRF4 DNA binding, leading to combined immunodeficiency with susceptibility to severe infections.ObjectivesWe aimed to develop an allele-specific editing strategy targeting the pathogenic IRF4 p.T95A variant and to evaluate genomic precision, safety, and functional rescue in edited T cells.MethodsBase and prime editing (PE) strategies were designed in a K562 reporter line, prioritizing allele-specific PE to minimize bystander and off-target effects. Patient-derived naïve T cells were selected and edited ex vivo via electroporation of PE and PE guide RNAs (pegRNAs). Functional rescue was assessed by flow cytometry and cytokine profiling of effector memory differentiation, Th17 polarization, and α4β7 expression. Genomic precision was evaluated with methods such as variant-aware in silico modeling and amplicon sequencing.ResultsThe proband is a 24-year-old man with a de novo IRF4 T95A variant, multidrug-resistant Mycobacterium avium colitis, recurrent candidiasis, and EBV-driven lymphoproliferation, for whom HSCT carries prohibitive risk. His T cells exhibit reduced effector memory differentiation, impaired Th17 responses, and diminished α4β7 expression. In the K562 reporter with the patient’s IRF4 allele, PE achieved efficient double-digit on-target correction with selective mutant-allele knockout while preserving the wild-type allele. Ongoing work focuses on optimizing PE in primary patient naïve T cells and evaluating phenotypic rescue and genome-wide specificity.ConclusionThis study demonstrates the feasibility of nonviral, allele-specific prime editing to correct a de novo IRF4 variant in autologous T cells, representing a promising personalized therapy for dominant IEIs.

CRISPR tiling screens have enabled the characterization of regulatory sequences but are limited by low resolution arising from the indirect readout of editing via guide RNA sequencing and enrichment analysis. This study introduces an end-to-end experimental assay and computational pipeline, which leverages targeted sequencing of CRISPR-introduced alleles at the endogenous target locus following dense base-editing mutagenesis. As a proof of concept, we studied a putative CD19 enhancer, an immunotherapy target in leukemia, and identified alleles and single nucleotides crucial for CD19 regulation. Our visualization tools revealed transcription factor motifs corresponding to the top-ranked nucleotides. Validation experiments confirmed that mutations in MYB, PAX5, and EBF1 binding sites reduce CD19 expression. Critically, editing MYB and PAX5 motifs conferred resistance to CD19 CAR-T cell therapy, revealing how non-coding variants can drive immunotherapy escape. Taken together, this approach achieves nucleotide-resolution genotype-phenotype mapping at regulatory elements beyond conventional gRNA-based screens.

ABSTRACT:Acute myeloid leukemia (AML) with TP53 mutations is almost universally refractory to chemotherapy, molecular-targeted therapies, and hematopoietic stem cell transplantation, leading to dismal clinical outcomes. The lack of effective treatments underscores the urgent need for novel therapeutic strategies. Using genome-wide CRISPR/Cas9 dropout screens in isogenic Trp53-wild-type (WT) and Trp53-knockout mouse AML models, combined with transcriptomic and proteomic analyses of AML samples from mice and humans, we identify the XPO7-NPAT (exportin 7-nuclear protein, coactivator of histone transcription) pathway as essential for TP53-mutated AML cell survival. In TP53-WT AML, XPO7 functions as a tumor suppressor by regulating the nuclear abundance of p53 protein, particularly when basal levels of functional p53 are high. However, in TP53-mutated AML, XPO7 drives leukemia proliferation by retaining NPAT, an XPO7-associated protein predominantly expressed in TP53-mutated AML, within the nucleus. NPAT depletion induces genome-wide histone loss, compromises genomic integrity, and triggers replication catastrophe in TP53-mutated AML cells. Notably, the analysis of publicly available AML data sets, primary AML samples, and single-cell intrapatient mRNA profiles further reveals elevated XPO7 and NPAT expression in TP53-mutated AML. Finally, we validate the XPO7-NPAT pathway as a critical driver of leukemia progression in vivo using patient-derived xenograft models of TP53-WT and TP53-mutant AML. Our study delineates key molecular mechanisms underlying TP53-mutated AML pathogenesis and identifies the XPO7-NPAT axis as a critical vulnerability in this refractory leukemia subtype.

Despite extensive investigation, the molecular control of developmental hemoglobin expression remains incompletely elucidated. Hemoglobin switching is controlled by transcription factors, miRNAs, and RNA-binding proteins (RBPs) that enforce gene regulatory changes through development. Here we examine the role of the heterochronically silenced N-6 methyladenosine (m6A) RNA-binding protein IGF2BP1 that was previously described to regulate HBG1/2 indirectly by suppressing BCL11A expression through an unknown mechanism. We find that IGF2BP1 binds and activates HIC2, itself a BCL11A repressor. Furthermore, we identify that IGF2BP1 plays a BCL11A-independent role by direct binding to HBG1/2 to promote its translation. Stop codon-proximal m6A-modified coding sequences within HBG2 transcripts are necessary and sufficient for direct positive regulation mediated by IGF2BP1. This work deepens the mechanistic understanding of hemoglobin switching and suggests a physical relationship between heterochronic RBPs and globin transcripts.

Long-read sequencing can characterize complex genome editing-induced DNA sequence changes such as large deletions, insertions, and inversions that are difficult to detect using short-read sequencing. However, PCR amplification and sequencing errors complicate accurate variant detection, and existing analysis tools are not optimized for gene editing specific allelic outcomes. Here we present CRISPRLungo, a computational pipeline specifically designed for long-read amplicon sequencing of gene edited samples. CRISPRLungo incorporates unique molecular identifier (UMI)-based error correction and statistical filtering to distinguish true editing events from background noise, enabling robust detection of small indels and structural variants. Through systematic benchmarking using simulated datasets, we demonstrate that CRISPRLungo outperforms existing approaches in both accuracy and read recovery. CRISPRLungo supports both Oxford Nanopore and PacBio platforms and identify previously undetected structural variant edits such as inversions in published CRISPR datasets. To demonstrate allele-specific edit quantification, we applied CRISPRLungo to analyze edited primary cells from a patient with harboring compound heterozygous SBDS mutations, accurately quantifying SBDS editing outcomes despite contaminating reads from the homologous SBDSP1 pseudogene. To maximize accessibility, we developed a fully client-side web application requiring no installation, making advanced long-read analysis accessible to researchers regardless of computational expertise. CRISPRLungo is freely available at https://github.com/pinellolab/CRISPRLungo with a user-friendly web interface available at https://pinellolab.github.io/CRISPRLungo .

Human-derived induced pluripotent stem cells (iPSCs) are an invaluable resource in both two-dimensional (2D) and three-dimensional (3D) tissue engineering due to their multilineage potential in culture systems. To date, modeling red blood cell (RBC) disorders such as sickle cell disease (SCD) from iPSCs has been challenging due to the tendency for differentiation protocols to produce immature erythrocytes that lack robust β-globin expression and enucleate poorly. Here, we demonstrated an optimized three-stage erythroid differentiation protocol that generates enucleated, β-globin-expressing RBCs from somatically sourced iPSCs, derived from both healthy donors and patients with homozygous SCD. Induced RBCs (iRBCs) present phenotypically as GlyA+Band3+CD71lo and express adult hemoglobin tetramers. SCD iRBCs displayed sickling phenotypes in vitro when exposed to hypoxia. RNA-sequencing analysis of iPSC-derived SCD reticulocytes revealed dysregulated disease-relevant molecular pathways, suggesting future therapeutic avenues of investigation can be identified in this model. We further refined the protocol into a xeno-free formulation by replacing albumin sources with polyvinyl alcohol (PVA), significantly enhancing iRBC production without loss of terminal maturation. The ability to generate patient-specific iRBCs from somatic cell sources provides a valuable in vitro tool for the study of SCD and the development of novel treatments.

Hematopoietic stem cells (HSCs) are self-renewing, multipotent, and engraftable precursors of all blood cells. Efficient delivery of therapeutic gene products and gene editing machinery to correct disease-causing gene variants in endogenous HSCs while they remain in the body holds exciting potential to leverage HSC potency for the treatment of monogenic blood disorders. Toward this goal, we used adeno-associated virus (AAV) to deliver CRISPR guide RNAs (gRNAs) to edit HSC genomes in situ in Ai9;SpCas9-EGFP transgenic mice carrying a Cas9-activatable Lox-STOP-Lox-tdTomato reporter cassette together with a constitutive SpCas9-2A-EGFP. Using a variety of conditions and vector designs, we tested whether systemic administration to these mice of AAVs carrying SpCas9compatible gRNAs designed to cut DNA upstream and downstream of the STOP cassette would induce tdTomato expression in HSCs. Our findings identify self-complementary AAVs (scAAVs) and increased ratio of guide to Cas9 as parameters facilitating higher editing efficiency. Of note, we find preserved multilineage output and engraftability of HSCs upon scAAV-gRNA editing. In an example application of this technology, we explore the potential for in situ HSC gene editing by dual AAV-CRISPR delivery and demonstrate robust gene modification, concurrent with induction of therapeutic fetal hemoglobin, in a sickle cell disease mouse model modified to express SpCas9. In summary, this work offers a sensitive and adaptable platform that allows robust modification of HSC genomes in situ.

Recently, cytosine base editors (CBEs) have emerged as a promising therapeutic tool for specific editing of single nucleotide variants and disrupting specific genes associated with disease. Despite this promise, the currently available CBEs have the significant liabilities of off-target and bystander editing activities, partly due to the mechanism by which they are delivered, causing limitations in their potential applications. In this study, we engineered optimized, soluble and stable Cas-embedded CBEs (CE_CBEs) that integrate several recent advances, which were efficiently formulated for direct delivery into cells as ribonucleoprotein (RNP) complexes. Our resulting CE_CBE RNP complexes efficiently target cytosines in TC dinucleotides with minimal off-target or bystander mutations. Delivery of additional uracil glycosylase inhibitor protein in trans further increased C-to-T editing efficiency and target purity in a dose-dependent manner, minimizing indel formation. A single electroporation was sufficient to effectively edit the therapeutically relevant locus BCL11A for sickle cell disease in hematopoietic stem and progenitor cells in a dose-dependent manner without cellular toxicity. Significantly, these CE_CBE RNPs permitted highly efficient editing and engraftment of transplanted cells in mice. Thus, our designed CBE proteins provide promising reagents for RNP-based editing at disease-related sites.

Therapeutic genome editing promises to transform medicine. Pivotal discoveries have provided a diverse and versatile set of tools to correct pathogenic mutations or produce protective alleles using CRISPR-based technologies. These innovative therapies are especially adaptable for blood and immune disorders, where clinical methods allow haematopoietic stem cells (HSCs) to be mobilized, harvested, engineered ex vivo and transplanted back into a patient to permanently replace their blood system. This paradigm has been exemplified with the first US Food and Drug Administration (FDA)-approved CRISPR-Cas9 therapy for sickle cell disease and β-thalassaemia, exa-cel (Casgevy). Although promising, efficient delivery of gene edits involves complicated ex vivo manipulation and toxic myeloablative conditioning. The quiescent and elusive nature of HSCs also brings associated challenges. In this Review, we explore the state-of-the-art genome editing technologies of nucleases, base editors and prime editors, which hold promise to address unmet clinical needs for patients with inherited haematological disorders. We highlight the progress made for several disorders and discuss the challenges that remain for ex vivo and in vivo targeting of HSCs for next-generation gene therapies.

Although therapeutic genome editing holds great potential to remedy diverse inherited and acquired disorders, targeted installation of medium to large sized genomic modifications in therapeutically relevant cells remains challenging. We have developed an approach that permits DNA sequence assembly and integration in human cells leveraging CRISPR-targeted dual flap synthesis. This method, named prime assembly, allows for RNA-programmable site-specific integration of single- or double-stranded DNA fragments. Unlike homology-directed repair, prime assembly was similarly active in dividing and non-dividing cells. We applied prime assembly to perform targeted exon recoding, transgene integration, and megabase-scale rearrangements, including at therapeutically relevant loci in primary human cells. Prime assembly expands the capabilities of genome engineering by enabling the targeted integration of medium to large sized DNA sequences without relying on double-stranded DNA donors, nuclease-driven double strand breaks, or cell cycle progression.

Targeted delivery of macromolecular therapeutics holds great promise for overcoming the limitations of conventional small molecules, enabling the modulation of protein-protein interactions and precise genome editing. However, efficient, safe, and cell type-specific delivery remains a major challenge. To address this, we developed a modular platform for synthesizing heterotrifunctional bio-orthogonal macromolecular conjugates (BMCs) by engineering diverse combinations of targeting ligands, cell-penetrating peptides (CPPs), and bioactive cargos. We optimized facile bioconjugation chemistries to generate BMCs with improved yields, structural integrity, and activity. Modular BMCs accommodate diverse components, including antibodies and receptor ligands for targeting, CPPs for intracellular trafficking, and optical probes, therapeutic peptidomimetics, and CRISPR-Cas9 nuclease as cargo to confer specific biological activities. We assayed their utility across multiple applications: BMCs with fluorescently labeled cargo revealed endosomal escape and intracellular accumulation; peptidomimetic MYB transcription factor inhibitor BMCs exhibited potent antileukemic activity against acute myeloid leukemia cells; and Cas9 BMCs achieved rapid delivery and cell type-specific gene editing in human cells. The BMC approach enables the customizable delivery of functional macromolecules, nominating BMCs as a broadly applicable platform for biomedical applications.

Sickle cell disease and β-thalassemia, two major β-hemoglobinopathies, pose significant clinical challenges globally. Current treatments often face limitations in efficacy and tolerability. The transcription factor ZBTB7A has emerged as a promising therapeutic target for reactivating fetal hemoglobin expression. Here, we report the discovery and characterization of SH6, a small molecule non-IMiD degrader of ZBTB7A. SH6 induces fetal hemoglobin in erythroid cell lines in a CRBN and ZBTB7A-dependent manner, and it is capable of inducing fetal hemoglobin expression in healthy donor, SCD and β-thalassemia patient CD34+ cell derived erythroid cells. The efficacy of SH6 is confirmed in a xenotransplantation humanized mouse model. SH6 outperforms currently available therapeutic agents in vitro , and shows synergy with hypomethylating agents. SH6 exhibits a favorable in vivo toxicity profile. Our findings establish SH6 as a promising therapeutic lead candidate for further optimization towards clinical development for treatment of sickle cell disease and β-thalassemia. ### Competing Interest Statement The authors have declared no competing interest. Cooley’s Anemia Foundation, https://ror.org/03hv05g61 United States Department of Defense, https://ror.org/0447fe631

Gene editing the BCL11A erythroid enhancer is a validated approach to fetal hemoglobin (HbF) induction for β-hemoglobinopathy therapy, though heterogeneity in edit allele distribution and HbF response may impact its safety and efficacy. Here, we compare combined CRISPR-Cas9 editing of the BCL11A +58 and +55 enhancers with leading gene modification approaches under clinical investigation. Dual targeting of the BCL11A +58 and +55 enhancers with 3xNLS-SpCas9 and two single guide RNAs (sgRNAs) resulted in superior HbF induction, including in sickle cell disease (SCD) patient xenografts, attributable to simultaneous disruption of core half E-box/GATA motifs at both enhancers. Unintended on-target outcomes of double-strand break (DSB) repair in hematopoietic stem and progenitor cells (HSPCs), such as long deletions and centromere-distal chromosome fragment loss, are a byproduct of cellular proliferation stimulated by ex vivo culture. Editing quiescent HSPCs bypasses long deletion and micronuclei formation and preserves efficient on-target editing and engraftment function.

The YAP-TEAD protein-protein interaction mediates YAP oncogenic functions downstream of the Hippo pathway. To date, available YAP-TEAD pharmacologic agents bind into the lipid pocket of TEAD, targeting the interaction indirectly via allosteric changes. However, the consequences of a direct pharmacological disruption of the interface between YAP and TEADs remain largely unexplored. Here, we present IAG933 and its analogs as potent first-in-class and selective disruptors of the YAP-TEAD protein-protein interaction with suitable properties to enter clinical trials. Pharmacologic abrogation of the interaction with all four TEAD paralogs resulted in YAP eviction from chromatin and reduced Hippo-mediated transcription and induction of cell death. In vivo, deep tumor regression was observed in Hippo-driven mesothelioma xenografts at tolerated doses in animal models as well as in Hippo-altered cancer models outside mesothelioma. Importantly this also extended to larger tumor indications, such as lung, pancreatic and colorectal cancer, in combination with RTK, KRAS-mutant selective and MAPK inhibitors, leading to more efficacious and durable responses. Clinical evaluation of IAG933 is underway. Chapeau et al. develop a nonallosteric inhibitor of the interaction between YAP and all four TEAD proteins. Treatment with the inhibitor, either as monotherapy or in combination with other treatment modalities, leads to induction of cell death in several in vivo cancer models.

Introduction. Chimeric antigen receptor T cells (CAR-T), bispecific and antibody-drug conjugates are promising adoptive immunotherapies that can overcome the limitations of conventional cancer treatments and have demonstrated striking efficacy when targeting dispensable lineage antigens (Ag), e.g. CD19 for B-ALL. Nonetheless, the absence of safely actionable tumor-restricted markers hampers their application to other hematological malignancies, such as acute myeloid leukemia (AML). Since AML shares most surface markers with normal hematopoietic stem/progenitor cells (HSPC) or differentiated myeloid cells, on-target/off-tumor toxicities would result in myeloid aplasia and impairment of hematopoietic reconstitution. Furthermore, due to AML intra-tumoral heterogeneity, targeting more than one Ag may be required, exacerbating the risk of overlapping toxicity. Despite this, AML immunotherapies are currently under development, but their role will likely be restricted to bridge treatment before allogeneic HSPC transplantation (HSCT), decreasing the chances of AML eradication. Removal of targeted Ag through CRISPR-Cas KO from donor HSPCs used in HSCT has recently been proposed, but this can only be applied to genes dispensable for hematopoietic function. However, targeting irrelevant genes may facilitate tumor escape through Ag downregulation. Here, we show that precise editing of the targeted epitope within FLT3, KIT (CD117) and IL3RA (CD123) in HSPCs results in loss of Ab binding without KO, preserving physiologic protein expression, regulation, and intracellular signaling. Critically, this strategy enables targeting one or more genes fundamental for leukemia survival, resulting in potent anti-leukemia efficacy with minimal on-target/off-tumor toxicity. Methods. Through epitope-mapping, we identified substitutions in the FLT3, KIT and CD123 extracellular-domains that avoid detection by therapeutic Abs. We validated the functionality of mutated receptors (ligand affinity, western-blot, proliferation, RNAseq, phospho-proteomic MS) and their resistance to on-target killing (mAb-affinity, CAR-T co-culture). We optimized a base-editing protocol to introduce these mutations in CD34+HSPCs and developed advanced in vivo models with co-engraftment of healthy HSPCs, patient-derived AML xenografts (PDX) and CAR-T to assess selective elimination of leukemia and protection of healthy hematopoiesis. Results. To develop our approach, we selected mAbs under development for AML therapy: clone 4G8 (FLT3), Fab-79D (KIT) and 7G3 (CD123). To identify residues involved in mAb binding, we designed Sleeping Beauty epitope mapping libraries. We found that single amino-acid substitutions can disrupt therapeutic mAb binding despite preserved surface receptor. Since epitope engineering can be achieved by point mutations, we reasoned that base editing (BE) could be a suitable and safer option compared to homology directed repair (HDR). By electroporating sgRNA+ABE variants, we achieved successful epitope editing with minimal toxicity. By using fluorescent FLT3L, SCF and IL-3, we confirmed preserved ligand binding to edited receptors. Activation of downstream signaling was confirmed by western blot. By performing in vitro killing assays, we found that, while cells expressing WT FLT3, KIT or CD123 were eliminated, those expressing epitope-edited variants were resistant to CAR-mediated killing and survived up to experiment termination without eliciting T cell activation and degranulation. To introduce our variants into human HSPCs, we optimized a BE protocol on mobilized peripheral blood-derived CD34+ cells, achieving up to 86%, 78% and 78% efficiency for FLT3, KIT and CD123, respectively. Contrary to previous observations with HDR editing, BE efficiencies were similar in bulk and primitive, HSC-enriched subsets (CD90+45RA-) with no skewing of stem phenotype. BE HSPCs were resistant to CAR or mAb-mediated killing in vitro. To confirm the safety of our approach, we compared FLT3, CD123, KIT edited to AAVS1 control HSPCs and found no differences in proliferative response, transcriptional changes (RNAseq), phospho-proteomic profile and colony-forming capacity. Xenotransplantation of BE HSPCs in NBSGW mice showed preserved repopulation and multilineage differentiation capacity, both in primary and secondary recipients. To assess if FLT3 CAR can eliminate AML while sparing FLT3-edited hematopoiesis, we sequentially engrafted NBSGW mice with HSPCs and human PDX cells. We observed a significant increase in the percentage of FLT3 edited cells in CAR-treated mice and relative depletion of CD19+ subsets (pre-B, pro-B), granulocytes, granulo-mono progenitors (GMP) and lymphoid-primed multipotent progenitors (LMPP) only in the AAVS1-BE group, while mice engrafted with FLT3-BE HPSC were protected. Concomitantly, mice treated with 4G8-CAR achieved complete AML eradication. CAR-T exposed to FLT3-BE hematopoiesis displayed lower PD-1 expression compared to AAVS1-BE. As done for FLT3, we transplanted CD123-BE HSPCs and confirmed multilineage repopulation comparable to controls. Mice treated with CD123 CAR-T showed eradication of AML cells and concomitant protection of epitope-edited myeloid lineages, including granulocytes, DCs and HSPCs. To test whether our approach allows multiplexing, we tested combinations of FLT3, KIT and CD123 editing, which provided additive protection from triple-specific CAR-T cells in vitro. Furthermore, combined dual FLT3+CD123 BE could protect hematopoietic lineages in vivo when mice were treated with FLT3+CD123 CAR-T, which in turn were able to eradicate PDXs resistant to FLT3-targeting alone. Discussion. Our studies provide proof of concept that tumor-associated Ags shared by normal tissue can be safely targeted by precisely modifying the epitope recognized by adoptive immunotherapies in healthy cells, endowing them with selective resistance and generating an artificial leukemia-restricted Ag. The innovative tools developed in this work can increase the therapeutic index of AML immunotherapies and enable long-term anti-leukemia maintenance. By restricting on-target activity to leukemia cells, epitope-editing can reduce the Ag burden to which CAR-T are exposed, decreasing undesired CAR-T stimulation, cytokine secretion and exhaustion. Epitope editing can easily be multiplexed to enable combination therapies while avoiding overlapping toxicities, further enhancing the chances for tumor eradication. Finally, epitope editing may be exploited to improve non-genotoxic conditioning for autologous gene therapy or HSCT, either alone or in combination with other therapeutic targets, to avoid depletion of transplanted cells and achieve in vivo selection of genome-engineered cells. Conclusion. We believe that epitope-engineering of HSPCs is a novel and highly promising technology that can enable safer and more effective immunotherapies when on-target/off-tumor toxicities are the key limiting factor to successful clinical translation. Citation Format: Gabriele Casirati, Andrea Cosentino, Adele Mucci, Mohammed S. Mahmoud, Iratxe Ugarte Zabala, Jing Zeng, Scott B. Ficarro, Denise Klatt, Christian Brendel, Alessandro Rambaldi, Jerome Ritz, Jarrod A. Marto, Danilo Pellin, Daniel E. Bauer, Scott A. Armstrong, Pietro Genovese. Epitope editing enables targeted immunotherapy of acute myeloid leukemia [abstract]. In: Proceedings of the American Association for Cancer Research Annual Meeting 2024; Part 2 (Late-Breaking, Clinical Trial, and Invited Abstracts); 2024 Apr 5-10; San Diego, CA. Philadelphia (PA): AACR; Cancer Res 2024;84(7_Suppl):Abstract nr NG05.

CRISPR tiling screens have advanced the identification and characterization of regulatory sequences but are limited by low resolution arising from the indirect readout of editing via guide RNA sequencing. This study introduces CRISPR-CLEAR, an end-to-end experimental assay and computational pipeline, which leverages targeted sequencing of CRISPR-introduced alleles at the endogenous target locus following dense base-editing mutagenesis. This approach enables the dissection of regulatory elements at nucleotide resolution, facilitating a direct assessment of genotype-phenotype effects.

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.

Despite over a century of work on hemoglobinopathies, a comprehensive understanding of hemoglobin switching and factors regulating hemoglobin expression remains incomplete even as the first CRISPR/Cas9 gene therapy has obtained approvals for the treatment of transfusion-dependent β-thalassemia and sickle cell disease. We studied factors related to the heterochronically silenced m6A RNA-binding protein IGF2BP1 which has been identified as a positive regulator of fetal hemoglobin expression by directly suppressing BCL11A expression. In contrast to this observation, post-transcriptional control by IGF2BP1 generally acts through improving the translation and mRNA stability of IGF2BP1 target mRNAs in an m6A-dependent manner. To better characterize the IGF2BP1-mediated mechanism responsible for control of fetal hemoglobin expression, we assessed whether BCL11A was required for IGF2BP1-mediated regulation of fetal hemoglobin and discovered a direct BCL11A-independent mechanism. Additionally, we characterize a heretofore poorly understood BCL11A-dependent mechanism for IGF2BP1 in regulating fetal hemoglobin expression. We developed a luminescent transgenic reporter to disentangle BCL11A-dependent and newly identified BCL11A-independent mechanisms and used this system to discover that IGF2BP1 directly supports the translation of HBG mRNA in a manner requiring intact stop codon-proximal m6A motifs which are absent from the HBB mRNA. Hemoglobin switching relies on a set of heterochronically regulated transcription factors, miRNAs, and RNA-binding proteins. This work identifies a novel mode of direct post-transcriptional regulation of HBG mRNA by a heterochronically regulated RNA-binding protein, IGF2BP1, that was previously described to regulate HBG indirectly via BCL11A. This work shows that HBG mRNA is bound directly by IGF2BP1 which supports HBG translation. HBB, not normally directly regulated by IGF2BP1, becomes responsive to IGF2BP1 overexpression upon addition of the HBG-derived stop codon-proximal m6A sites to an HBB transgene, a strategy with potential to be exploited for the improved expression of globin transgenes for hemoglobinopathy gene therapy.