Genome editing of HBG1 and HBG2 to induce fetal hemoglobin

Sickle cell disease (SCD) and β-thalassemia are common disorders caused by HBB gene mutations that alter quantity or quality of the β-globin subunit of adult hemoglobin (HbA, α2β2).^1,2  Severely affected individuals experience multiorgan damage, with substantial morbidity and early mortality. Αllogeneic hematopoietic stem cell (HSCs) transplantation can be curative but carries high risk of severe toxicities, particularly for patients who lack fully histocompatible donors.³  Hence, new methods for autologous gene therapy are being sought.

Genome editing of patient HSCs by clustered regularly interspaced short palindromic repeats (CRISPR)–Cas9 nucleases represents a promising approach for genetic correction of β-hemoglobinopathies.^4-6  These nucleases introduce targeted DNA double-stranded breaks (DSBs) that can be exploited therapeutically through 2 general cellular DNA damage repair strategies. First, HBB mutations can be corrected via homology-directed repair (HDR).^5,7-12  Second, fetal hemoglobin (HbF, α2γ2) can be induced in adult red blood cells (RBCs) by using nonhomologous end-joining (NHEJ) mediated mutations to disrupt noncoding DNA regulatory elements that repress transcription of the genes encoding γ-globin (HBG1 and HBG2) postnatally.^4,13-19  Numerous clinical studies show that elevated HbF production is associated with reduced morbidity and mortality in SCD and β-thalassemia.^20,21  In extreme cases, a rare, benign genetic condition called hereditary persistence of HbF (HPFH) induces high-level pancellular HbF expression in RBCs and eliminates the pathologies of coinherited β-hemoglobinopathies.²²  Therapeutically, the induction of HbF by NHEJ may offer several advantages over direct HBB repair for treating β-hemoglobinopathies. First, NHEJ is the dominant DNA DSB repair pathway and is active in all phases of the cell cycle, which is particularly relevant to editing quiescent HSCs. Second, correction of the SCD mutation via HDR is accompanied by undesired NHEJ-mediated insertion/deletion (indel) mutations in cis or trans.^7,23  Third, in contrast to HDR, NHEJ does not require delivery of an exogenous DNA donor template, which introduces additional technical challenges in manufacturing. Lastly, genome editing to induce HbF provides a single generally applicable therapeutic strategy for many different β-hemoglobinopathy mutations, whereas correction by HDR would require mutation-specific optimization.

Several HPFH mutations occur in a region 118 to 114 nt upstream of the HBG1 and HBG2 transcription start sites and disrupt a cognate-binding element for the γ-globin gene repressor BCL11A (TGACC).^24,25  Previously, we targeted this region in CD34⁺ hematopoietic stem and progenitor cells (HSPCs) by lentiviral expression of Cas9 and associated single guide RNAs (sgRNAs) followed by in vitro differentiation.¹⁶  The percentage of HbF (%HbF) was increased to potentially therapeutic levels in the RBC progeny of most CD34⁺ cells with on-target edits. Here we advance that proof-of-concept study by achieving several essential requirements for clinical translation, including transient Cas9:sgRNA delivery to HSPCs, high-level editing in human HSCs capable of multilineage engraftment after transplantation into immunodeficient mice, and absence of detectable off-target mutations or deleterious hematopoietic effects. Therefore, Cas9 ribonucleoprotein (RNP)–mediated disruption of the BCL11A repressor binding site in the promoters of HBG1 and HBG2 is a potentially feasible and safe therapeutic strategy for treating SCD and β-thalassemia.

Plerixafor-mobilized CD34⁺ cells from patients with SCD were collected according to the protocol Peripheral Blood Stem Cell Collection for Sickle Cell Disease Patients (www.clinicaltrials.gov identifier #NCT03226691), which was approved by the human subject research institutional review boards at the National Institutes of Health and St. Jude Children’s Research Hospital. All patients provided informed consent.

Mice were housed and handled in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. Animal experiments were carried out in accordance with a protocol (Genetic Tools for the Study of Hematopoiesis) approved by the institutional animal care and use committee of the St. Jude Children’s Research Hospital or Boston Children’s Hospital.

The antibodies used in this study are listed in supplemental Table 1. The cytokines used are listed in supplemental Table 2. The oligonucleotides used are listed in supplemental Table 3. The isolation, editing, and analysis of CD34⁺ cells before and after xenotransplantation are described in the supplemental Methods.

We edited human CD34⁺ HSPCs by electroporating RNP complexes of Cas9:sgRNA-1 targeting the BCL11A consensus motif at position −118 to −114 in the HBG1/HBG2 gene promoters (Figure 1A). Initial studies were performed using the Neon Transfection System (see supplemental Methods). We titrated Cas9 and sgRNA-1 with or without 2′-O-methyl modifications and 3′-phosphorothioate internucleotide linkages shown previously to improve gene editing rates in primary cells²⁶  and electroporated the RNP complexes into adult human peripheral blood–mobilized CD34⁺ HSPCs. Transfected cells were cultured for 4 days and then analyzed for small indels by next-generation sequencing (NGS) of a 318-bp polymerase chain reaction (PCR) fragment centered around the RNP cleavage site (Figure 1B; supplemental Figure 1A-B). The maximum indel frequencies observed were ∼45% with unmodified sgRNA and 80% with chemically modified sgRNA. After electroporation, cell viability declined with increasing RNP concentration but remained >80% under all conditions (supplemental Figure 1C-E). Importantly, the 9 most frequently observed indel alleles, and presumably most others, disrupt the BCL11A-binding site and therefore are predicted to derepress the associated γ-globin genes (Figure 1C).

We cultured gene-edited CD34⁺ HSPCs from 2 donors under erythroid differentiation conditions and measured the HbF expression in late-stage erythroblasts. The mean indel frequency measured was 63% ± 18% (Figure 2A). As usual for in vitro–derived erythroblasts, background HbF expression was high; ∼50% of the cells stained with HbF antibody (termed F-cells), and HbF protein constituted ~10% of the total Hb in cell lysates (Figure 2B-C; supplemental Figure 2). Nonetheless, gene editing of the HBG1 and HBG2 gene promoters raised %F-cells and %HbF protein significantly. Editing did not alter the expression of erythroid maturation markers Band3 or CD49d (Figure 2D) or the conversion of nucleated erythroblasts to anucleate reticulocytes (Figure 2E).

To assess our capacity to edit repopulating HSCs, we electroporated CD34⁺ cells with Cas9:sgRNA-1 RNP and transplanted them into nonobese diabetic/severe combined immunodeficiency/Il2rγ^−/−/Kit^W41/W41 (NBSGW) mice, an immunodeficient strain that supports the development of human erythroid precursors without conditioning.²⁷  We measured the fraction of engrafted human cells and their indel frequencies serially in blood and in bone marrow after the mice were euthanized at 17 weeks, when donor CD34⁺ cells present derive mainly from HSCs.²⁷  No significant differences in chimerism occurred between edited and nonedited donor-derived CD45⁺ hematopoietic cell populations (Figure 3A). Chimerism levels were similar for edited and nonedited donor T cells (hCD45⁺/CD3⁺), B cells (hCD45⁺/CD19⁺), myeloid cells (hCD45⁺/CD33⁺), and erythroid cells (CD45⁻/CD235a⁺) in bone marrow at 17 weeks (Figure 3B).

The indel frequencies of edited human donor cells declined from 70.9% ± 5.8% a few days after editing to 41.8% ± 9.4% at 17 weeks after xenotransplantation (Figure 3C), similar to what has been reported previously.^14,23,28  Notably, the 13-nt HPFH mutation occurred at similar frequencies in CD34⁺ HSPCs before and after transplantation (Figure 3C; supplemental Figure 3). Indel frequencies were also similar in CD34⁺ donor cell–derived myeloid, erythroid, and B cells (Figure 3D), indicating that editing did not alter the development of these lineages from HSCs. The indel spectrum in edited cells was similar before and after transplantation, with no clonal dominance in the latter populations (Figure 3E; supplemental Figure 3).

Circulating human RBCs are short lived and difficult to detect in mouse xenotransplantation studies. Therefore, we studied immune-selected CD235a⁺ donor-derived erythroblasts from the bone marrow of recipient mice at 17 weeks after transplantation. We obtained mainly late-stage hemoglobinized erythroblasts and reticulocytes, with no obvious differences in morphology, expression of maturation markers, or enucleation fraction between the edited and control populations (Figure 3F-H). Notably, background %HbF was lower in control (nonedited) erythroblasts generated in vivo compared with in vitro cultures (compare Figure 2B-C to Figure 3I-J). The percentages of both HbF-immunostaining cells (F cells) and HbF protein compared with overall Hb were significantly increased in edited vs control erythroid cells generated in vivo (Figure 3I-J). Considering that the F-cell fraction was 51.3% ± 12.5% in erythroblasts with ∼42% indels (Figure 3D), it is likely that HbF was induced in all edited cells, thereby favoring therapeutic efficacy.²⁹  The %HbF in erythroid cell lysates correlated with indel frequency (Figure 3K). These xenotransplantation studies demonstrate that it is feasible to induce erythroid HbF expression by Cas9 RNP-mediated disruption of the HBG1/HBG2 BCL11A-binding motif, with no obvious alterations in hematopoietic development in vivo.

The tandem HBG1 and HBG2 genes harbor nearly identical nucleotide sequences, including recognition sites for sgRNA-1. Simultaneous RNP-induced DSBs at both genes can result in the deletion of the intervening 4.9-kb region,^16,30  leaving a single hybrid gene with HBG2 promoter sequences fused to the downstream HBG1 gene (Figure 4A). We developed 2 assays as proxies for the 4.9-kb deletion. First, we used a TaqMan quantitative PCR assay to quantify loss of DNA in the HBG2-HBG1 intergenic region ≥366 bp upstream of the RNP cleavage site (“Δ366⁺”). Second, we PCR-amplified exon 3 of HBG1 and HBG2, using a common primer pair, and performed NGS to quantify loss of HBG2 (“ΔHBG2”) based on a single-nucleotide difference between HBG2 and HBG1 (G vs C at cDNA position 410) (Figure 4A). Both assays showed that the 4.9-kb deletion rates were ∼30% before transplantation and 11% at 17 weeks posttransplantation (Figure 4B). By comparison, small indel rates were ∼72% and 43% before and after transplantation, respectively.

To measure large indels clonally and investigate whether the associated loss of HBG2 impaired HbF production, we analyzed burst-forming unit erythroid (BFU-E) colonies from edited and control CD34⁺ cells (Figure 4C). The frequency of hematopoietic colonies was decreased by ∼30% in CD34⁺ cells electroporated with Cas9:sgRNA-1 RNP vs Cas9 alone (supplemental Figure 4A), with the proportions of myeloid and erythroid colonies being similar (supplemental Figure 4B). In pools of ∼1000 gene-edited colonies, the small indel frequency was 57.3% (supplemental Figure 4C) and the %HbF increased from 8.1% ± 0.6% in controls to 27.7% ± 2.5% after editing (supplemental Figure 4D). In 124 BFU-E colonies analyzed individually, the frequency of small on-target indels was 45%, and the frequency of large indels measured by loss of HBG2 exon 3 was 24% (Figure 4C). Colonies with no on-target edits contained 9.5% ± 5.8% HbF (n = 47). Most edited colonies contained either 4 γ-globin alleles, all with small indels that disrupted BCL11A binding motifs (n = 15; HbF, 48.4% ± 30%), or 1 deleted HBG2 allele with the BCL11A-binding motifs disrupted in the 3 remaining γ-globin genes (n = 28; HbF, 40.8% ± 17%). Thus, the loss of a single HBG2 allele, presumably reflecting the 4.9-kb deletion, did not appreciably alter the %HbF.

Simultaneous dsDNA breaks at the HBG1 and HBG2 promoters can also cause inversion of the intervening DNA (Figure 4D), predicted to eliminate the expression of both genes. Using primer pairs that detect this inversion, we identified multiple PCR products specifically in edited HUDEP-2 cells and CD34⁺ HSPCs. Sanger sequencing showed that these inverted segments contained interrupted sequences and deletions ranging from 236 to 1916 bp. We detected the inversion in 1 of 96 BFU-E colonies (1%) generated from bulk-edited CD34⁺ HSPCs with an overall on-target small indel frequency of 84%. Thus, this inversion likely occurs at low rates.

We characterized the genome-wide activity of Cas9:sgRNA-1 by incubating RNP with purified genomic DNA followed by circularization for in vitro reporting of cleavage effects by sequencing (CIRCLE-seq), a sensitive in vitro method for enriching for Cas9-cleaved genomic DNA and identifying the breakpoints by NGS.³¹  In 3 experimental replicates, we consistently detected 26 potential off-target candidate sites (Figure 5, left panel). However, no mutations were detected at these sites by NGS of PCR products generated from edited HSPCs (sensitivity 0.1%-0.01%) (Figure 5, right panel).

At ≥16 weeks after xenotransplantation, gene-edited human donor cells contained no chromosomal rearrangements detected by G-band karyotyping (supplemental Figure 5) or fluorescence in situ hybridization (FISH) with a probe located distal to the γ-globin gene loci on human chromosome 11 (supplemental Figure 6). Overall, our data suggest that Cas9:sgRNA-1 RNP targets the HBG1 and HBG2 promoters with high specificity and without detectable off-target DNA mutations at the sites analyzed.

We next analyzed the effects of Cas9:sgRNA-1 gene editing in CD34⁺ HSPCs from 2 individuals with SCD. As granulocyte colony-stimulating factor (G-CSF) is contraindicated in SCD,^32,33  we edited plerixafor-mobilized peripheral blood CD34⁺ cells. After xenotransplantation into NBSGW mice, edited and control (nonedited) CD34⁺ cells populated the bone marrow similarly (Figure 6A) and gave rise to similar fractions of human T, B, myeloid, and erythroid cells (Figure 6B). For SCD donor 1, the small on-target indel frequency was 79% before transplantation and remained high at 16 weeks after transplantation (72% ± 3%) (Figure 6C). For SCD donor 2, the indel frequency was 55% before transplantation and declined to 18% ± 2% after transplantation. The spectrum of on-target indels in SCD CD34⁺ HSPCs was similar to that observed after editing normal G-CSF–mobilized CD34⁺ HSPCs and did not change appreciably after xenotransplantation (supplemental Figure 7).

The frequency of 4.9-kb HBG2-HBG1 intergenic deletions in edited SCD donor 1 CD34⁺ cells detected by the ΔHBG2 and Δ366⁺ assays (Figure 4A) was ∼30% before transplantation and 33% after transplantation (Figure 6D). These large deletions were less frequent in edited SCD donor 2 CD34⁺ cells, consistent with relatively low frequencies of small indels. At 16 weeks after xenotransplantation with edited SCD donor 1 HSPCs, we purified donor CD34⁺ cells and analyzed them by FISH using a probe for the HBG2-HBG1 intergenic region and a control probe for an adjacent downstream region including the HBB gene (supplemental Figure 8A). Most cells derived from gene-edited donor HSPCs (42 of 75 cells analyzed) were heterozygous for the 4.9-kb HBG2-HBG1 intergenic deletion (overall allele frequency, 32%) (supplemental Figure 8B).

At 16 weeks after transplantation, bone marrow erythroblasts derived from gene-edited and control CD34⁺ HSPCs exhibited similar morphologies (Figure 6E) and maturation profiles as determined by flow cytometry (supplemental Figure 9A-B). Erythroblasts derived from gene-edited HSPCs from SCD patient 1 CD34⁺ cells contained 77.8% ± 1.9% F cells as compared with 15.7% ± 1.6% in control nonedited erythroblasts (Figure 6F; supplemental Figure 9C). The %HbF was 42.9% ± 1.5% after gene editing vs 1.3% ± 0.5% in control erythroblasts (Figure 6G). Erythroblasts generated in vivo from HSPCs from SCD patient 2 also exhibited substantially increased HbF induction after undergoing gene editing, despite lower on-target indel frequencies (Figure 6C,F-G). HbF induction in SCD erythroid cells correlated with indel formation (Figure 6H), similar to that which we observed in erythroid cells derived from normal donor HSPCs (Figure 3K). To investigate the potential therapeutic benefits of HbF induction, we used immunomagnetic beads to purify CD235a⁺ erythroid cells from the bone marrow of mice that had been transplanted with HSPCs from SCD patient 1. After being subjected to hypoxia (2% O₂) for 8 hours, 32% ± 1.0% of control erythroid cells exhibited sickled morphology as compared with 13.5% ± 1.0% of gene-edited cells (Figure 6I-J). Thus, HbF induction resulting from Cas9:sgRNA-induced disruption of the BCL11A-binding site of the HBG1 and HBG2 gene promoters inhibits erythroid cell sickling.

Our findings demonstrate that genome editing of the HBG1 and HBG2 promoters in HSCs induces HbF in erythroid progeny. However, on-target indel frequencies varied between CD34⁺ cell donors and tended to decline over time after xenotransplantation (Figures 3C and 6C). The Streptococcus pyogenes Cas9 protein used in our studies described thus far contains 2 tandem C-terminal SV40 nuclear-localization signals (NLSs; Cas9-2xNLS).³⁴  We recently reported improved gene editing of human HSPCs using a modified SpCas9 containing a c-Myc–like NLS at the N terminus and both SV40 and nucleoplasmin NLSs at the C terminus (Cas9-3xNLS) (Figure 7A).³⁵ 

To test whether Cas9-3xNLS edits HSPCs more consistently, we electroporated RNP complexes containing sgRNA-1 and either Cas9-2xNLS or Cas9-3xNLS into normal human CD34⁺ cells with the Lonza Nucleofector 4-D platform and then measured the resulting indel frequencies after 3 days of culture in HSC media. Cas9-3xNLS maintained consistently high indel rates (65.9% to 84.2%) across 2 donors and 4 different delivery conditions, whereas under identical conditions, Cas9-2xNLS/sgRNA-1 RNP generated more variable indel rates, ranging from 24.1% to 82.1% (Figure 7B).

Next, we transfected Cas9-3xNLS/sgRNA-1 RNP into CD34⁺ HSPCs from 2 normal donors and transplanted the cells into NBSGW mice. At 16 weeks after transplantation, human hematopoietic chimerism in the bone marrow measured by human CD45 expression was 93.6% ± 0.9% for donor 1 and 56.5% ± 5.8% for donor 2 (Figure 7C). This difference may arise from variation in the fraction of repopulating HSCs within the bulk donor CD34⁺ cell population from different donors. Bone marrow chimerism for T, B, myeloid, and erythroid cell populations was similar for edited and control donor cells, although erythroid chimerism was low for donor 2 (Figure 7D). The indel frequencies measured by TIDE (tracking of indels by decomposition³⁶ ) of donor CD34⁺ HSPCs were similar before and after transplantation, ranging from 86% to 98%. The frequency of the 4.9-kb intergenic HBG2-HBG1 deletion in CD34⁺ donor cells at 16 weeks after transplantation estimated by the TaqMan Δ366⁺ assay was ∼40% for donor 1 and 22% for donor 2 (Figure 7E). We purified donor-derived CD34⁺ cells from recipient bone marrow and cultured them under erythroid differentiation conditions. The HbF level was 25% ± 5% in RBCs derived from gene-edited donor CD34⁺ cells but only 6% ± 1% in control RBCs (Figure 7F). Together, the results of these studies indicate that CD34⁺ HSPCs edited with Cas9-3xNLS/sgRNA-1 RNP maintained consistently high levels of editing after xenotransplantation. Although additional comparisons are required, our experiments suggest that optimized delivery of RNP with Cas9-3xNLS or similar enhanced-activity variants should facilitate robust editing of the HBG1 and HBG2 promoters in long-term engrafting HSCs, with HbF being induced in the RBC progeny to therapeutic levels for SCD and β-thalassemia.

Previously, we demonstrated that Cas9-mediated disruption of a repressor element in the HBG1 and HBG2 promoters induces erythroid HbF expression.¹⁶  Here, we advance those results by showing that transient expression of Cas9 RNPs in human CD34⁺ cells results in high-frequency editing of the target site (≤80%) in repopulating HSCs with induction of HbF to potentially therapeutic levels in RBC progeny and no detectable genotoxicity. Editing at this site precisely disrupts a binding motif for the BCL11A transcriptional repressor protein, consistent with findings that most on-target indels activate HBG1/HBG2 transcription.¹⁶  Our study generated several important new findings.

First, our methods achieved substantially higher rates of target-site editing in HSCs than reported previously after transient expression of Cas9 RNPs³⁰  or transcription activator-like effector nucleases,³⁷  which will likely translate to greater therapeutic benefits. The variability of HSC editing that we observed in our studies using Cas9-2xNLS could be due to differences in donors, CD34⁺ mobilization methods (ie, G-CSF vs plerixafor), or CD34⁺ cell purification. Notably, the use of chemically modified sgRNA and Cas9-3xNLS³⁵  achieved consistently high editing activity across donors and delivery methods, in contrast to a commonly used Cas9 with 2 SV40 NLSs fused to the C terminus (Cas9-2xNLS). The improvement in editing by Cas9-3xNLS may arise from NLS signals at both ends of the protein and/or the use of 3 unique NLS signals that interact synergistically with nuclear import machinery.³⁸ 

Second, our study demonstrates high-frequency editing (≤70%) of plerixafor-mobilized CD34⁺ HSPCs from individuals with SCD, which represents the cell product most likely to be manipulated therapeutically. Increased HbF levels enhance the survival of erythroblasts and mature RBCs in patients with β-thalassemia or SCD.^39-46  The therapeutic benefits of our strategy depend on the fraction of repopulating HSCs edited and the resultant levels and cellular distribution of HbF. Because circulating human RBCs are cleared very rapidly in mice, we are unable to measure the survival advantage conferred by gene editing. After allogeneic bone marrow transplantation for SCD, ∼20% donor HSC chimerism can result in 100% circulating donor RBCs and marked clinical improvement.^47,48  In natural history studies, pancellular HbF levels >20% to 30% are suggested to protect against SCD morbidities, consistent with biochemical studies of HbS polymerization.^49-51  Based on these parameters extrapolated from clinical studies, the preclinical outcomes achieved in this study (70% to 80% indel rate in HSCs, with most or all RBC progeny expressing HbF at an average level of 30% to 40%) are likely to produce clinical benefits. One limitation of our study is that on-target editing frequencies in HSCs varied considerably between experiments and donors in initial experiments. However, our data using newly developed Cas9-3xNLS suggest that consistent high-level editing of the HBG1/HBG2 promoters in repopulating HSCs can now be achieved, similar to what was recently reported for an erythroid-specific enhancer in the BCL11A gene.³⁵ 

Third, our study provides critical new preclinical safety assessments by showing that the 4.9-kb deletion resulting from simultaneous on-target DSBs in HBG1 and HBG2 promoters is not deleterious to HSCs or RBC precursors. Analysis of clonal BFU-E colonies showed that HbF expression was not impaired by deletion of HBG2. In most clones harboring the 4.9-kb deletion, the BCL11A-binding motif was destroyed at the DSB repair site, thereby favoring expression of the reconstituted HBG1 gene. Expression of β-like globin genes is facilitated via interaction with an upstream enhancer, termed locus control region.⁵²  Loss of HBG2 may eliminate competition for the locus control region, thereby favoring upregulation of the remaining HBG1 gene. Interestingly, a germline variant similar to the 4.9-kb deletion noted in our study has been identified in multiple unrelated individuals. This variant arises from unequal crossing over between microsatellite markers in the second introns of HBG2 and HBG1, resulting in a 5-kb deletion that generates a single chimeric γ-globin gene with exons 1 and 2 originating from HBG2 and exon 3 from HBG1.^53,54  Individuals with this variant have RBCs with reduced Gγ to Aγ ratio as infants but are otherwise normal throughout life. These clinical observations, along with our experimental data, suggest that the 4.9-kb deletion arising from our gene-editing strategy will not have deleterious effects on globin expression, erythroid development, or RBC physiology.

Additionally, our studies did not detect off-target DSBs in edited HSCs at top candidate sites identified by CIRCLE-seq, and no chromosomal abnormalities were detected by G-banding or FISH analysis of >200 CD34⁺ HSPCs after xenotransplantation. For ultimate therapeutic application, it will be important to examine all sites reproducibly identified by CIRCLE-seq after editing the HBG promoters with Cas9-3xNLS and perform further assessments for structural variants induced by our gene-editing protocol.

In summary, our studies demonstrate that it is possible to achieve high-level editing of the HBG1/HBG2 promoters in G-CSF–mobilized CD34⁺ cells from normal individuals and plerixafor-mobilized CD34⁺ cells from individuals with SCD, that editing does not impair hematopoiesis, and that productive editing results in HbF induction to potentially therapeutic levels, with no off-target DSBs or chromosomal rearrangements being detected. These preclinical safety and efficacy studies support a promising strategy for modifying human HSPCs to treat human β-hemoglobinopathies.

The authors thank the following core facilities and individuals at St. Jude Children’s Research Hospital: Flow Cytometry (Richard Ashmun, Deanna Langfitt, and Stacie Woolard), Animal Resource Center (Chandra Savage), and Veterinary Pathology (Patricia J. Varner).

This work was supported by National Institutes of Health, National Heart, Lung, and Blood Institute grant P01 HL053749 (D.E.B., S.P.-M., S.Q.T., and M.J.W.), Doris Duke Charitable Foundation grant DDCF-2017093 (S.Q.T. and M.J.W.), National Institutes of Health, National Heart, Lung, and Blood Institute grant U01HL145793 (S.Q.T.), National Institute of Allergy and Infectious Diseases R01AI117839, and National Institute of General Medical Sciences R01GM115911 (S.A.W.), The Assisi Foundation of Memphis (M.J.W.), and St. Jude/ALSAC and the St. Jude Collaborative Research Consortium, “Novel Gene Therapies for Sickle Cell Disease." The St. Jude Cytogenetic Shared Resource Laboratory is supported by National Institutes of Health, National Cancer Institute grant P30 CA21765 and American Lebanese Syrian Associated Charities.

Contribution: J.-Y.M. contributed to study design, experiments, analyses, and drafting of the manuscript; P.A.D. contributed to experiments, analyses, and drafting of the manuscript; T.M. and V.K. contributed to experiments, data analysis, and drafting of manuscript; S.C.F. contributed to experiments and mouse maintenance; M.M.H., K.L., M.D.N., N.U., and S.A.W. contributed reagents; S.K., S.N.P., S.S.P., D.S., K.J.W., J.Z., Y.W., and Y.Y. contributed to experiments; C.R.L. contributed to experiments and data analysis; S.T.P. contributed to data analysis; B.Y.R., A.S., and J.F.T. contributed to study design and contributed reagents; D.E.B. and S.P.-M. contributed to study design; and S.Q.T. and M.J.W. contributed to study design, data analysis, and drafting of the manuscript.

Conflict-of-interest disclosure: M.J.W. is a consultant for GlaxoSmithKline, Rubius Inc., Cellarity Inc., Beam Therapeutics, and Esperion; none of the consulting work is relevant to the current project. The remaining authors declare no competing financial interests.

Correspondence: Mitchell J. Weiss, Department of Hematology, St. Jude Children’s Research Hospital, 262 Danny Thomas Pl, Memphis, TN 38105; e-mail: mitch.weiss@stjude.org; and Shengdar Q. Tsai, Department of Hematology, St. Jude Children’s Research Hospital, 262 Danny Thomas Pl, Memphis, TN 38105; e-mail: shengdar.tsai@stjude.org.