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Enhanced fetal hemoglobin production via dual-beneficial mutation editing of the HBG promoter in hematopoietic stem and progenitor cells for β-hemoglobinopathies

Stem Cell Research & Therapy volume 15, Article number: 504 (2024) Cite this article

Abstract

Background

Sickle cell disease (SCD) and β-thalassemia patients with elevated gamma globin (HBG1/G2) levels exhibit mild or no symptoms. To recapitulate this natural phenomenon, the most coveted gene therapy approach is to edit the regulatory sequences of HBG1/G2 to reactivate them. By editing more than one regulatory sequence in the HBG promoter, the production of fetal hemoglobin (HbF) can be significantly increased. However, achieving this goal requires precise nucleotide conversions in hematopoietic stem and progenitor cells (HSPCs) at therapeutic efficiency, which remains a challenge.

Methods

We employed Cas9 RNP-ssODN-mediated homology-directed repair (HDR) gene editing to mimic two naturally occurring HBG promoter point mutations; -175T > C, associated with high HbF levels, and −158 C > T, a common polymorphism in the Indian population that induces HbF under erythropoietic stress, in HSPCs.

Results

Asymmetric, nontarget ssODN induced high rates of complete HDR conversions, with at least 15% of HSPCs exhibiting both the −175T > C and −158 C > T mutations. Optimized conditions and treatment with the small molecule AZD-7648 increased this rate, with up to 57% of long-term engrafting human HSPCs in NBSGW mice containing at least one beneficial mutation. Functionally, in vivo erythroblasts exhibited high levels of HbF, which was sufficient to reverse the cellular phenotype of β-thalassemia. Further support through bone marrow MSC co-culture boosted complete HDR conversion rates to exceed 80%, with minimal InDels, improved cell viability, and induced fetal hemoglobin levels similar to those of Cas9 RNP-mediated indels at BCL11A enhancer and HBG promoter.

Conclusions

Cas9 RNP-ssODN-based nucleotide conversion at the HBG promoter offers a promising gene therapy approach to ameliorate the phenotypes of β-thalassemia and SCD. The developed approach can simplify and broaden applications that require the cointroduction of multiple nucleotide modifications in HSPCs.

Background

β-Hemoglobinopathies, encompassing sickle cell disease (SCD) [1] and β-thalassemia [2], represent the most commonly inherited monogenic disorders worldwide [3], posing a substantial public health challenge. These disorders are characterized by mutations in the HBB gene locus, leading to defective β-globin production that can result in vaso-occlusive crisis and hemolysis in SCD or ineffective erythropoiesis in β-thalassemia [1, 2]. Epidemiological studies estimate that there are more than 300,000 new cases of β-hemoglobinopathies each year, with SCD accounting for the majority of cases. The burden of these diseases is particularly heavy in low- and middle-income countries [3], including India, where they impose significant socioeconomic costs. Consequently, there is an urgent need for universal, curative treatments.

Recent therapeutic advancements have made the prospect of curing β-hemoglobinopathies a reality [4]. Gene therapy has emerged as a promising alternative to allogeneic hematopoietic stem cell transplantation (HSCT). Modifying patient hematopoietic stem and progenitor cells (HSPCs) to express a recombinant form of hemoglobin (βA−T87Q), encoded by a self-inactivating BB305 vector, has shown clinical efficacy in both β-thalassemia [5] and SCD [6]. The FDA recently approved this therapy, which is now commercially known as Zynteglo and Lyfgenia, for β-thalassemia and SCD, respectively. Furthermore, progress in gene editing technologies has expanded therapeutic gene manipulation strategies, expanding the genetic medicine toolbox for β-globin defects [4, 7]. Among these, Casgevy, which employs CRISPR/Cas9 to reactivate fetal hemoglobin (HbF) by targeting the erythroid-specific enhancer of BCL11A, has been commercially approved [8, 9].

The therapeutic landscape is expanding with strategies aimed at either correcting HBB locus mutations directly [10,11,12] or inducing the expression of developmentally silenced fetal hemoglobin from the HBG locus [13,14,15]. The erythroid-specific downregulation of BCL11A is a leading approach for HbF reactivation [4, 9]. This is closely followed by point/deletional mutations in the HBG promoter, which have been shown to effectively derepress γ-globin in cellular and animal models [13,14,15,16,17].

The −175T > C mutation in the HBG promoter is a high-potency hereditary persistence of fetal hemoglobin (HPFH) point mutation that creates a new enhancer motif (CAGATG) adjacent to the GATA1 binding site [18, 19]. This mutation facilitates the formation of a protein complex of GATA1, TAL1, LIM domain only 2 (LMO2) and LIM domain binding protein 1 (LDB1) that activates γ-globin expression by mediating a long-range interaction of the gamma-globin promoter with its distal enhancer LCR [20]. Clinical data suggest that the −175T > C HPFH mutation results in significant increases in Aγ and Gγ globin levels of up to ~ 40% and ~ 30%, respectively, making it a target for therapeutic genome editing [19, 21, 22]. Recent studies have attempted to introduce this mutation into HSPCs using Cas9 RNP-adeno-associated virus (AAV6) and base editing technologies, achieving 3.48% and 39% efficiency, respectively, in long-term-engrafting human HSPCs [14, 16].

Another genetic variant, the −158 C > T Xmn1 polymorphism (HGB2 rs7482144, ‘Senegal’ and ‘Arab-Indian’ haplotypes), is associated with milder disease phenotypes on coinheritance with β-hemoglobinopathies [23,24,25]. This polymorphism has not been previously targeted for HBG promoter editing but is known to elevate γ-globin levels under erythropoietic stress, a hallmark of β-hemoglobinopathies [26]. At least two studies have reported individuals with both −175T > C and − 158 C > T mutations who exhibited normal hematologic indices and pancellular distribution of HbF in red blood cells, despite having HbS mutation [26, 27]. There are no studies thus far to mimic both of these beneficial mutations in patient HSPCs for therapeutic application.

In this study, we employed a Cas9 ribonucleoprotein (RNP)- and single-stranded oligonucleotide (ssODN)-based homology-directed repair (HDR) strategy to simultaneously introduce the −175T > C and −158 C > T mutations into the HBG promoter in HSPCs. We achieved an ex vivo editing efficiency of up to 80% and an in vivo efficiency of up to 57%, successfully reactivating γ-globin expression to correct the cellular phenotype of β-hemoglobinopathies.

Materials and methods

CD34 + HSPCs purification and culture

Mobilized peripheral blood (mPB) in the leftover blood bags post allogenic transplantation was collected at Christian Medical College Hospital, Vellore, following the approval of the appropriate Institutional Review Board (IRB). Mononuclear cells were isolated through density gradient centrifugation utilizing Lymphoprep™ (#07801), and CD34 + cells were subsequently selected via immunomagnetic separation employing the EasySep™ Human CD34 Positive Selection Kit II (Stem Cell Technologies, Cat# 17856). CD34 + cells were cultured at a density of 2 × 105 cells/ml in StemSpan™ SFEM II medium supplemented with SCF (240 ng/ml), Flt3 (240 ng/ml), TPO (80 ng/ml), IL6 (40 ng/ml), and 1X antibiotic-antimycotic, as described in our earlier studies [28, 29]. Additionally, a cocktail comprising small molecules—resveratrol, UM729, and SR1—collectively referred to as RUS—was added to the culture medium in the specified experiments [30].

Gene editing reagents

sgRNAs targeting the HBG1/2 promoters were manually designed. An sgRNA against the control editing target AAVS1 was designed using the Synthego CRISPR design tool. The synthetic guide RNAs were purchased as lyophilized products from Synthego (Menlo Park, CA, USA). The ssODNs were purchased from Integrated DNA Technologies (IDT) with custom backbone PS modifications. Recombinant WT Cas9 was purchased from Takara Bio (Cat no. 632640).

Gene editing and erythroid differentiation of HSPCs

A total of 2 × 105 HSPCs were suspended in nucleofection solution and combined with 25 pmols RNP and 80 pmols of ssODN. The editing mix was loaded onto the wells of the electroporation unit and processed with Amaxa™ 4D-Nucleofector™ using the pulse code DZ-100. After nucleofection, the cells were briefly incubated with 100 µl of prewarmed media and plated in culture media.

Alternately, cells were suspended in HyClone™ MaxCyte® electroporation buffer, followed by RNP ssODN addition at the same dose, and loaded onto the processing assembly (OC25 × 3 or OC100 × 2) for electroporation with pulse code HSC-5 in the MaxCyte® ExPERT® GTx™ unit. The processing unit was immediately placed directly in a 37 °C CO2 incubator and plated in culture media. HSPCs were differentiated in vitro into erythroblasts as described in our previous work [29]. The functional analysis of erythroblasts, such as flow cytometry analysis and globin chain and hemoglobin variant analyses, was also performed as described in our previous study [29].

For screening experiments to test HDR enhancers, HSPCs were treated with small molecules (Supplementary Table 1) for 24 h post electroporation. The absolute HDR-positive cell counts were determined by multiplying HDR editing frequency with cell counts at the time of collection.

For co-culture experiments, MSCs were seeded in required cell culture dishes at a density of 10,000 cells/cm2 in MSC expansion media (MEM-Alpha modification with 10% FBS and 1x antibiotic), a day prior to seeding edited HSPCs in MSC. The MSC media is removed gently, followed by 2 washes with 1X PBS, to remove remnants of serum. Edited HSPCs are then seeded over the MSCs in HSPCs expansion media supplemented with AZD-7648.

Genotyping analysis

Genomic DNA extraction was performed 3–5 days post nucleofection utilizing QuickExtract™ DNA Extraction Solution. Over 10,000 cells were collected and rinsed with 1X PBS. The cell pellet was then resuspended in approximately 20–50 µl of QuickExtract™ solution and subjected to sequential incubation at 68 °C and 98 °C. The extracted DNA was directly processed for amplification of the target using the appropriate primers (Supplementary Table 2). Following the confirmation of bands through gel electrophoresis, the amplified product was purified using the Macherey-Nagel™ NucleoSpin™ PCR Clean-up Kit. For Sanger sequencing, the purified PCR amplicon was processed with BigDye™ Terminator v3.1 Cycle Sequencing, and the resulting chromatograms were analyzed by the Synthego ICE or TIDE algorithm. In the case of next-generation sequencing (NGS), the purified PCR amplicon is subjected to nested PCR for the ligation of adapters. The amplicons were then purified using the MN Nucleospin PCR Clean-up Kit and processed for indexing and library preparation, followed by sequencing on the Illumina platform. The resulting reads were analyzed using CRISPResso2 software.

Xenograft assay

Experiments involving murine subjects were conducted in accordance with the guidelines set forth by the Institute Animal Ethics Committee. Cells collected in PBS approximately 24 h after nucleofection were infused into in-house bred female NOD. Cg-KitW-41JTyr+PrkdcscidIl2rgtm1Wjl/ThomJ (NBSGW) mice [31], preconditioned with busulfan (12.5 mg/kg) 48 h prior, via tail vein injection. The age of the mice ranged from 6 to 8 weeks. Analysis of engraftment and human chimerism was carried out 16–18 weeks after transplantation, employing the antibody combinations specified in our previous work [30]. Cells positive for hCD235a were sorted using BD FACS ARIA™ III and subsequently processed for F-cell evaluation via intracellular staining and flow cytometry.

All the other methods are detailed in the supplementary methods.

The work has been reported in line with the ARRIVE guidelines 2.0.

Results

Asymmetric, nontarget ssODN introduces high-frequency HDR conversions at the HBG promoter in HSPCs

An ssODN-based templated correction strategy for HDR was utilized to introduce beneficial point mutations to activate the HBG promoter. The ssODN donor template contained 3 nucleotide changes, namely, -175T > C, -158C > T, and −153G > C. The −153G > C mutation served as a PAM shield to prevent Cas9 recutting. While the −158C > T position was located close to the Cas9 cut site (1st nucleotide downstream), the −175T > C was positioned 17 nucleotides upstream (Fig. 1A). It has been reported that the position of the desired nucleotide for modification, the target locus, and the design of ssODNs can significantly impact HDR conversion [32]. A screening of ssODN designs against the target was performed directly in the HSPCs. This screen included 8 designs with varying features, such as length, symmetry with respect to the cut site, and polarity against the two strands. Designs B1-B4 were non-target, whereas designs C1-C4 were target strand sequences (Fig. 1B). The HSPCs were edited with Cas9-RNPs and each designed ssODNs (Fig. 1C). An analysis using CRISPResso of deep sequencing reads from the edited cells indicated that the approach resulted in complete HDR conversions with all 3 intended nucleotide conversions at the HBG promoter, albeit at varying rates for different donor designs (Fig. 1D, and E). The B-series ssODNs with nontarget strand sequences achieved better complete HDR than the C-series ssODNs with target strand sequences. Specifically, compared with other designs, symmetric ssODN (B2) and asymmetric ssODN with longer 3’ ends (B4) showed high complete HDR conversion rates of approximately 15%. Other events that arose due to incomplete HDR conversions, such as single conversions at the − 175th and − 158th positions and combinations of these with each other and PAM, were also beneficial for HbF induction. Compared with the B2 design, the asymmetric, nontarget ssODN B4 displayed increased total HDR rates (30% of HSPCs with beneficial mutations—either single or combinations of 2 or 3 mutations) (Fig. 1F and Supp Fig. 1A). All C-series ssODNs showed incomplete conversions, demonstrating the importance of ssODN design screening with respect to the position of the nucleotide to be gene edited.

Fig. 1
figure 1

Asymmetric, nontarget ssODN contributes to improved HDR at HBG promoter in HSPCs. (A) Schematic view of the target depicting the β-like globin gene cluster, encompassing the locus control region (LCR) (gray bars), followed by the HBE1 (gray box), HBG2 and HBG1 (red boxes), HBD (pink box) and HBB (beige box) genes sequentially from the 5’ to 3’ direction. Expanded view of HBG1/2 promoters detailing the gRNA target sequence (underlined in gray), protospacer adjacent motif (underlined in green), and the position of intended nucleotide conversions, -175T > C, -158 C > T and − 153G > C (PAM shield), are indicated. (Image made with GraphPad-Prism). (B) Graphic representation of ssODN oligo designs screened for HDR editing of HBG1/2 promoters. The target/gRNA binding strand is depicted in blue, and the nontarget/nongRNA binding strand, in green. The ssODNs - B series (green) and C series (blue), are colored according to their homologous derivative strand. Three desired mutations are marked as colored circles: -175T > C (maroon), -158 C > T (red), and − 153G > C (yellow). The dotted line denotes the Cas9 cut site, and the length scale is provided in base pairs. (C) Schematic workflow describing the steps for screening and validation of ssODNs (1B) for HDR editing of HSPCs. (Created with BioRender.com). (D) Representative NGS reads from CRISPresso2 output of edited samples showing alleles around the cut site. The three desired nucleotide conversions are indicated in colors (as described in 1 A, 1B). (E) The complete HDR frequency, i.e., the fraction of reads with conversions of all 3 nucleotides resulting from screened ssODN designs obtained from deep sequencing reads analyzed using CRISPResso2 (results displayed with the mean ± SEM for n = 2). (F) Deep sequencing results of edited samples showing total cumulative frequencies of all beneficial HDR conversions at the HBG1/2 promoters, analyzed using CRISPResso2 (n = 2; n = 1 for C2 due to low cell yield post editing in one replicate). (mean ± SEM)

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HBG promoters with − 175T > C and − 158 C > T reactivate HbF in differentiated erythroblasts

The phenotypic effects of the conversions generated in the HBG promoter are found within the erythroid lineage cells that arise from edited HSPCs. Consequently, the erythroid effects of editing with the designs that generated high HBG promoter conversions (B2 and B4) were compared against those of RNP (w/o ssODN). A three-phase erythroid differentiation protocol was applied to the HSPCs [33], and the differentiated cells were analyzed for downstream effects of editing (Fig. 2A). On analyszing the HbF expression using intracellular staining followed by flow cytometry both B2 and B4 displayed significantly greater HbF levels (~ 60%) than the unedited control (~ 18%) (Fig. 2B and C).

In the reverse-phase-HPLC chain analysis, the chromatogram confirmed the elevation of gamma-globin levels in B2 and B4 samples compared to the control. An increased height of both the Aγ and Gγ peaks was observed, suggesting simultaneous editing at both promoters, although it was greater for Aγ (Fig. 2D and E). RNP editing did not contribute to a remarkable increase in gamma globin chains. A 1.8-fold increase in the absolute number of HDR edited cells was produced by B4 ssODN compared with B2, suggesting that B4-ssODN has better viability than B2; thus, subsequent experiments were performed with B4-ssODN (Supp. Figure 1B). To confirm that the nucleotides converted in the HSPCs was retained in the differentiated cells, the B4-ssODN-edited HSPCs were plated on Methocult medium, and individual colonies were picked and sequenced. After the analysis of a total of 26 colonies, the complete HDR rates of the colonies were found to be similar to those of the seeded HSPCs (Fig. 2F).

The generation of γ-globin chains is attributed to their ability to form functional HbF tetramers (α2γ2). An increase in HbF tetramers was detected in B4, contributing to 22.00% of the total hemoglobin variants (Fig. 2G and H). Erythroid maturation of edited cells was similar to that of the control (Supp Fig. 1C and D).

To understand the contribution of each mutation to the modulation of HbF levels, the HSPCs were edited with ssODNs containing either − 175T > C or -158 C > T or both − 175T > C and − 158 C > T (B4). All these ssODNs contain a PAM shield mutation, -153G > C. To determine whether the PAM shield − 153G > C adversely affects HbF production, we also tested ssODNs with − 175T > C and − 158 C > T and without a PAM. HbF analysis revealed that the − 175T > C mutation is a major contributor to HbF, and the − 158 C > T mutation moderately influences HbF under normal erythropoiesis conditions (high editing efficiency in comparison with − 175T > C but relatively low HbF levels) (Fig. 2I). This finding is consistent with the role of the − 158 C > T mutation in reactivating γ-globin only under erythropoietic stress, such as β-thalassemia and SCD. The analysis also indicated that the PAM shield mutation does not affect HbF levels, as similar levels of HbF were observed in B4 and with the − 175T > C and − 158 C > T mutations.

Fig. 2
figure 2

HBG promoters with − 175T > C and − 158 C > T dual conversions expresses HbF in in vitro differentiated erythroblasts. (A) Schematic of the workflow followed for erythroid differentiation and analysis of edited cells (Created with BioRender). (B) Representative flow cytometry plots for globin expression analysis by intracellular HbF staining of differentiated erythroblasts in vitro. (C) Percentage of HbF+ ve mature erythroblasts observed on day 20 of erythroid differentiation from n = 3 samples (mean ± SEM; ****p < 0.0001, *** p < 0.001; one-way ANOVA (Dunnett’s multiple comparisons test)). (D) RP-HPLC analysis of traces of globin chains detected in erythroid protein lysates of control and edited samples on day 20 of differentiation. The areas under the Aγ and Gγ peaks are shaded in red. (E) Percentage of total γ globin chains (Aγ + Gγ) detected by RP-HPLC in control and edited samples. n = 3 to 5 (mean ± SEM; ** p < 0.01; one-way ANOVA (Dunnett’s multiple comparisons test). (F) Complete HDR frequency (reads containing all 3 mutations) in the edited HSPC pool and the CFU colonies generated from the HSPC edited pool. The HDR editing was analysed by Sanger sequencing, followed by Synthego ICE knock-in analysis. n = 26 CFU colonies. (G) Representative HPLC traces for hemoglobin tetramer variant analysis of protein lysates from day 20 erythroblasts. The areas under HbF and HbA are shaded red and gray, respectively. (H) Proportion of hemoglobin (Hb) tetramers detected by HPLC analysis (mean ± SEM for n = 3; ** p < 0.01; unpaired t-test). (I) Comparison of HDR conversion frequency at each desired nucleotide location with respect to HbF levels detected by flow cytometry in edited samples (mean ± SEM for n = 2)

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The − 175T > C and − 158 C > T HDR edits occur in primitive HSCs with a 2-fold reduced 4.9-kb deletion

To enhance the efficiency of HDR gene editing and the viability of HSPCs, we tested ssODNs modified with 1, 2, or 3 terminal phosphorothioate (PS) at both ends or 2 PS at only the 3’ or 5’ end of the B4 ssODN (Supplementary Fig. 2A) [34]. Compared with unmodified ssODN (B4-P0), PS-modified ssODNs resulted in a greater number of HDR-edited HSPCs. Specifically, the greatest number of HDR-edited HSPCs were generated by ssODNs with 2 PS modifications only at the 3’ end (Supplementary Fig. 2B). Consequently, B4-ssODNs with 2-PS modifications at the 3’ end were used in all subsequent experiments.

Previously, a cocktail of small molecules, Resveratrol, SR-1, and UM729 (RUS), was reported by us to enhance Cas9-RNP gene editing in the primitive subpopulation when it was added to the HSPCs culture medium [30]. When the RUS-supplemented HSPC culture was tested for RNP-ssODN editing, a modest increase of 10% in the complete HDR rate was observed in the total cells (Supplementary Fig. 2C). Notably, complete HDR events occurred in primitive CD34 + CD90 + HSCs, almost at a similar frequency as in the CD34 + CD90- progenitor population (Supplementary Fig. 2D), suggesting long term repopulating stem cells are also getting edited. Next, we examined the 4.9 kb deletions associated with HBG1/G2 editing and observed that RNP editing is associated with a 4.9 kb deletion at a frequency of 46% of cells, which is reduced by 2-fold in B4-edited HSPCs (Supplementary Fig. 2E).

Furthermore, the kinetics of HDR editing were examined by collecting HSPCs at various time points postelectroporation. Interestingly, HDR events were detected as early as 6 h postelectroporation and peaked at 12 h. No further increase in HDR events was observed after 12 h in the HSPCs from the two different donors (Supplementary Fig. 2F and 2G).

The DNA-PK inhibitor AZD-7648 enhances the effectiveness of ssODN-mediated complete HDR conversion in HSPCs

To further increase the efficiency of HDR mediated gene editing in HSPCs, complete HDR conversion was monitored in the presence of 24 selected small molecules known to regulate various cellular processes, such as the cell cycle, DNA repair, chromatin state, and immune responses [35,36,37,38,39,40]. HSPCs were electroporated with RNP and ssODN and then treated with selected small molecules for a brief window of 24 h. After withdrawal of the small molecules, the HSPCs were further cultured for 48 h with cytokines and collected for genotyping (Fig. 3A). The screen identified NU7441, and entinostat as the top candidates that influenced the complete HDR conversion, with mean efficiencies of 44, and 34%, respectively, over that of the DMSO control (10%) (Fig. 3B).

Given that NU-7441, an inhibitor of DNA protein kinase (DNA-PK) [41], promoted the highest HDR in the initial analysis, we focused on DNA-PK inhibition, and questioned whether any other DNA-PK inhibitor could outperform NU-7441. Consequently, the influence of various DNA-PK inhibitors—NU7441, AZD-7648 [42], DNMB, LTURM 34, and NU-7026—was analyzed. Among the candidates tested, AZD-7648 outperformed NU7441 and other DNA-PK inhibitors in elevating HDR (67.5 ± 8.51%) from that of the DMSO Control (16.25 ± 2.78%). Significantly, the frequency of NHEJ reads in the edited pool was substantially decreased to near-zero levels following AZD-7648 treatment (Fig. 3C) from a previous high frequency of 58.75 ± 3.6% without small molecule addition (DMSO).

Although AZD-7648 treatment induced very high HDR conversions and prevented InDels at a dose of 20 µM, cellular toxicity was also exhibited, as indicated by the reduced cell numbers at 72 h post-treatment (Fig. 3D). Reducing the dose to 7 μm, improved viability; however, it also reduced the frequency of HDR conversion. When the HSPCs were cultured with the RUS cocktail, the 7 μm dose produced complete HDR rates similar to that of the 20 μm dose of standard conditions (Fig. 3E and F). A treatment duration of 24 h was critical to obtain high HDR conversion (Supplementary Fig. 3A and B). Hence, a working dose of 7 μm and a 24-hour treatment window postelectroporation with AZD-7648 were used for further experiments. Treatment did not compromise the subpopulation composition of HSPCs or the proportion of colony-forming units (Supplementary Fig. 3C and D), although it reduced the counts (Fig. 3G).

Fig. 3
figure 3

Transient DNA-PK inhibition using AZD-7648 amplifies HDR editing in HSPCs. (A) Schematic of the experimental workflow for small molecule screening to enhance HBG promoter-targeted HDR conversion in HSPCs. (Created with BioRender.com). (B) Complete HDR conversion (reads with all 3 mutations) frequency in edited HSPCs treated with indicated small molecules and genotyped on day 3 postelectroporation by Sanger sequencing and Synthego ICE knock-in analysis (mean ± SEM for n = 4). (C) Complete HDR, NHEJ, and WT read frequencies observed on day 3 postelectroporation after transient exposure to various DNA-PK inhibitors (mean ± SEM for n = 4; **p < 0.01; two-way ANOVA (Dunnett’s multiple comparison test)). (D) HSPCs cell numbers after transient treatment with the DNA-PK inhibitors NU-7441 and AZD-7648 and the DMSO control on day 3 postelectroporation (mean ± SEM for n = 4; ****p < 0.0001; one-way ANOVA (Dunnett’s multiple comparison test)). (E) The frequency of complete HDR after treatment with different doses (0, 7, or 20 µM) of AZD-7648 with and without RUS supplementation in HSPC culture media. The data were analyzed using Sanger sequencing and ICE knock-in. The baseline HDR from the vehicle control is marked with a dotted line. (Mean ± SEM for n = 4; * p < 0.05; unpaired student t-test). (F) Cell numbers observed after treatment with different concentrations (0, 7, or 20 µM) of AZD-7648 with or without RUS supplementation on day 3 postelectroporation (mean + SEM for n = 2; ** p < 0.01; ordinary one-way ANOVA (Dunnett’s multiple comparison test)). (G) No. of CFU-E-, BFU-E-, CFU-GM- and CFU-GEMM colonies formed from HSPCs that are cultured with RUS and treated with DMSO or 7 µM AZD-7648- for 24 h post gene editing with Cas9 RNP and B4-ssODN (mean + SEM for n = 2; * p < 0.05; unpaired student t-test)

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Long-term engrafted HSPCs retain HDR editing and produce HbF-expressing erythroid cells in vivo

Analysis of the functional effects of gene editing in an in vivo model can validate the therapeutic efficacy of the strategy. HSPCs that had undergone electroporation for AAVS1 control, RNP-B4 ssODN (B4), and RNP-B4 ssODN along with AZD-7648 treatment (B4-AZD-7648) were infused into busulfan-conditioned NBSGW mice via the tail vein (Fig. 4A). The Maxcyte GTx electroporator was utilized to prepare the infusion cell products for the animal experiments as it allows easy scale-up.

At 16 weeks post-transplantation, all animals were sacrificed, and tissues were collected from the homogenized cell suspensions for analysis. Analysis of human CD45 + cells chimerism in the peripheral blood, bone marrow, and spleen revealed a similar level of engraftment for the three conditions tested: AAVS1, B4, and B4-AZD-7648 (Fig. 4B). No significant changes were observed in the 16-week hematopoietic output from the engrafted HSPCs within the bone marrow under any of the three conditions when the proportions of B cells (CD19+), myeloid cells (CD13+), T cells (CD3+), and erythroid cells (CD235+) were examined by flow cytometry (Fig. 4C, Supplementary Fig. 4A and B). HbF + ve cells were analyzed within the FACS-purified CD235+ ve erythroid cells, which exhibited a 3-fold increase after B4 editing and a 5-fold increase after B4-AZD-7648 editing (Fig. 4D and E). Genotyping of the engrafted cells was performed to verify that HbF activation occurred through the − 175T > C and − 158 C > T edits. The HDR efficiency was found to vary among the animals, consistent with earlier observations [12]. Animals infused with AZD-7648-treated edited HSPCs had at least 20% HDR edits in vivo, and the highest HDR-edited cell engraftment was 57%, among which 35% of the cells had complete HDR (Fig. 4F). To further support these findings, the engrafted bone marrow cells were subjected to in vitro erythropoiesis, and HPLC chain analysis confirmed HbF activation in the B4 and B4-AZD-7648 groups, with a significant increase in the therapeutic γ/β-like ratio in the B4-AZD-7648 group (Fig. 4G and H). The 4.9 kb deletions were also maintained in vivo with AZD-7648 treatment, resulting in up to 45% deletions (Supplementary Fig. 4c).

Fig. 4
figure 4

HDR-edited HSPCs engraft long-term in NBSGW mice and recapitulate HbF in vivo. (A) Schematic of the experimental plan for xenotransplantation assay of -175T > C- and − 158 C > T-HDR-edited HSPCs into NBSGW mice and subsequent analysis. (Created with BioRender.com). (B) The frequency of human cell (hCD45+) engraftment in the peripheral blood, spleen and bone marrow was detected at 16 weeks post infusion by flow cytometry. (n = 3–4; mean ± SEM). (C) Sixteen-week multilineage repopulation output of edited cells detected by staining for B cells (CD19), myeloid cells (CD33), and T cells (CD3) gated within the hCD45 + population and erythroid cells (CD235a) gated directly from the bone marrow P1 population (n = 3–4; mean ± SEM). (D) Representative flow cytometry zebra plots for intracellular HbF staining of sorted CD235a + erythroblasts in 16th week bone marrow. (E) Frequency of HbF+ ve cells among CD235a + erythroblasts sorted from bone marrow. (n = 3–4; mean ± SEM; *p < 0.05, ** p < 0.01; ordinary one-way ANOVA (Dunnett’s multiple comparison test)). (F) Total beneficial HDR frequencies at HBG1/2 promoters consisting of complete HDR (reads with all 3 conversions) and individual − 158 C > T conversions detected by genotyping 16th week bone marrow cells. The data are shown for individual animals analyzed in each edited group. (G) Reverse-phase high-performance liquid chromatography (HPLC) trace files of protein lysates from 16 th week NBSGW bone marrow cells differentiated in vitro. A representative chromatogram of a single animal from each treatment group is shown. The Gγ and Aγ peaks are shaded in red. (H) Globin chain ratio of β-like globins in in vitro differentiated long term engrafted bone marrow cells [γ/(γ + β)] (mean ± SEM for n = 3–4 technical replicates for 1 biological replicate; ** p < 0.01; ordinary one-way ANOVA (Dunnett’s multiple comparison test))

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The − 175T > C and − 158 C > T edits efficiently reverse the β-thalassemia phenotype

Next, the contribution of -158 C > T in alleviating the disease phenotype was investigated. As the effect of -158 C > T conversions can be studied only under a disease background, the initiation codon ATG > ACG β-thalassemia mutation was introduced in healthy donor HSPCs (wild type) through base editing to model β-thalassemia (Fig. 5A) [43]. This approach avoids the collection of HSPCs from β-thalassemia patients and provides wild-type HSPCs as an isogenic control to understand the efficiency of disease reversal by various gene editing approaches. The base-edited HSPCs contained 74% of the desired ATG > ACG start codon β-thal mutation. An equally efficient bystander mutation, which should not affect the thal phenotype as it is after the start codon mutation, was also present (Fig. 5B and C).

The β-thalassemia-modeled HSPCs were further edited with RNP and − 158 C > T ssODN or RNP and B4 ssODN with all 3 (-175T > C, -158 C > T and PAM shield) mutations. The complete HDR efficiencies were 40% and 44%, respectively (Fig. 5D). During in vitro erythropoiesis, a strong erythroid maturation defect was shown by β-thal-modeled HSPCs compared with wild-type HSPC-derived cells (reticulocytes: 30.2% vs. 53% in wild-type cells). The reticulocyte count was improved to 45% and 44% by the − 158 C > T and − 175T > C conversions, respectively (Fig. 5E). HbF levels were also high in the HDR-edited samples (Fig. 5F), although the basal HbF levels were also elevated in the control thalassemia modelled cells, reflecting the phenotype observed in patients. Similarly, ROS levels per cell in β-thalassemia-modeled erythroblasts were reduced by 50% in both the − 158 C > T and − 175T > C edited samples (Fig. 5G). All these experiments demonstrated that both mutations are beneficial for reversing the β-thalassemia phenotype, with − 158T > C being particularly beneficial under disease background.

Fig. 5
figure 5

Dual-beneficial conversions at HBG promoters reverses the phenotype of in vitro generated β-thalassemia model. (A) Schematic of the experimental workflow for generating thalassemia-modeled HSPCs (β-thal-HSPCs) via ABE base editing and further reversal of the phenotype via HBG1/2 editing. β-thal-HSPCs are generated by targeting the HBB initiation codon. The ABE base edited cells were subsequently electroporated with Cas9 RNP and ssODN. The cells were further differentiated into erythroid lineages for downstream analysis (Created with BioRender.com). (B) Frequency of base edits at the HBB gRNA target site. The intended ATG target and bystander T > C conversions were obtained by genotyping the DNA collected on day 10 post RNP + B4 electroporation via Sanger sequencing and analyzing the ab1 reads via EditR software (n = 1). (C) EditR sequence analysis output of the base-edited sample. The locations of T nucleotide conversions are highlighted by red dotted lines. The heatmap shows the observed nucleotide frequencies of all 4 bases spanning the 20-base pair long sgRNA target. (D) HDR frequency for base-edited samples electroporated with − 158-PAM and B4 (all 3 mutations) ssODNs. (E) Fraction of erythroid subsets observed on the 20th day of differentiation analyzed by flow cytometry using CD235a and CD71 staining. Populations were distinguished as reticulocytes (CD235a + CD71-), erythroblasts (CD235a + CD71+), erythroid progenitors (CD235a-CD71+) and undifferentiated cells (CD235a-CD71-) (n = 2 technical replicates analyzed per treatment). (F) Flow cytometry analysis of F + ve cell staining in day 20-derived erythroblasts from unedited control, β-thal-HSPC control and HDR-edited TM-HSPCs (n = 2 technical replicates analyzed per treatment). (G) The mean fluorescence intensity of DCFDA + staining was used to detect reactive oxygen species (ROS) in erythroblasts on the 12th day of differentiation. (n = 2 technical replicates analyzed per treatment)

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MSc coculture mitigates the toxicity associated with HDR gene editing in HSPCs

We hypothesized that the efficiency of RNP-ssODN mediated HDR gene editing could be further improved if the toxicity associated with RNP-ssODN and AZD-7648 treatment was alleviated. Recently, it was shown that anti-inflammatory factors produced by bone marrow MSCs overcome the toxicity associated with RNP-AAV6 HDR editing, resulting in better engraftment [44]. We questioned whether the same phenomenon applies to HSPCs edited with RNP-ssODN.

The HSPCs were electroporated with RNP-ssODN and were immediately either plated on MSC stromal cells or cultured under the regular protocol (Standard condition-SC). The influence of two sources of MSCs, bone marrow (BM) and Wharton’s jelly (WJ), was tested in the presence or absence of AZD-7648 (Fig. 6A, Supplementary Fig. 5A and 5B). Seventy-two hours post electroporation, the frequency of complete HDR improved from 14% in standard conditions to 22% and from 57% in the AZD-7648 group to 84% in the BM-MSC group. WJ-MSCs had similar effects on HDR but to a lesser extent than BM-MSCs (Fig. 6B). The absolute number of HDR-positive cells, a measure of viable HDR-edited cells, was improved by 7.8-fold in the BM-MSCs group treated with AZD-7648. A lesser effect was observed in WJ-MSCs (Fig. 6C). Similarly, the number of colonies produced in the metho-cult medium under standard and AZD-7648 conditions was increased by BM-MSCs co-culture (Fig. 6D). The proportion of colony output did not change under any of these conditions (Fig. 6E). The co-culture with BM-MSCs effectively maintained the proportion of primitive HSCs (Supplementary Fig. 6A) and reduced 4.9 kb homologous promoter deletion, which was elevated after AZD-7648 treatment (Supplementary Fig. 6B).

Gene editing approaches using Cas9 RNP have demonstrated remarkable efficiency in generating InDels, reaching over 90%, and have gained commercial approval [8, 9]. We compared our optimized HDR gene editing conditions with the Cas9 RNP-mediated InDels at the BCL11A enhancer and BCL11A binding region in the HBG promoter(-115 HBG promoter) (Fig. 7A). The results showed that our optimized conditions produced similar outcomes in terms of CFU colony proportions (Fig. 7B) and erythroid differentiation (Fig. 7C) when compared to cells edited at the BC11A enhancer and − 115 HBG promoter. Notably, the percentage of HbF+ ve cells (Fig. 7D and E), globin chains (Fig. 7F), and HbF tetramers (Fig. 7G and H) were comparable across these different edited conditions. These findings suggest that our HDR gene editing approach can achieve similar levels of effectiveness in both editing efficiency and functional outcomes as established NHEJ-based methods. All these experiments highlight the benefits of ssODN-mediated gene editing and culturing HSPCs with BM-MSCs to mitigate the toxicity associated with RNP-ssODN editing and AZD-7648 treatment (Fig. 8).

Fig. 6
figure 6

Coculture of electroporated HSPCs with MSCs mitigates HDR editing-associated toxicity. (A) Schematic of the experimental workflow for evaluating the efficacy of postelectroporation coculture of HSPCs with bone marrow and Wharton’s jelly derived mesenchymal stem cells (MSCs) in preventing cell toxicity. (Created with BioRender.com). (B) Frequency of reads corresponding to complete HDR (all 3 conversions), NHEJ indels, and wild-type sequences in edited samples genotyped by Sanger sequencing and Synthego ICE knock-in analysis. The standard condition (SC) refers to only cytokine culture (details in methods). Mesenchymal stem cells (MSCs) are derived from either bone marrow (BM-MSCs) or Wharton’s jelly (WJ-MSCs). All conditions were tested with and without AZD-7648. BM-MSCs and WJ-MSCs were also cultured in HSPC media supplemented with cytokines, both with and without AZD-7648. (mean ± SEM for n = 5; ** p < 0.01; unpaired student t-test). (C) Absolute number of complete HDR edited cells on day 3 postelectroporation. The fold increase for samples with respect to the control (SC) is indicated above the respective bars. (D) The number of colonies with different CFUs from 200 cells seeded in Methocult medium for 14 days was determined by microscopy (n = 3 per sample for 2 treatments performed per condition). (E) Proportion of CFU colonies obtained for different treatments on the 14th day. (Data are shown for n = 3 per sample for 2 treatments performed per condition.)

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Fig. 7
figure 7

HBG promoter with − 175T > C and − 158 C > T mutations generate functional outcome comparable to existing Indel-based gene editing approaches for HbF reactivation. (A) Genotype analysis to evaluate complete HDR (reads with all 3 mutations) and indel frequency in electroporated HSPCs by Sanger sequencing and Synthego ICE knock-in analysis (mean + SEM for n = 2). (B) Proportion of CFU-E-, BFU-E-, CFU-GM- and CFU-GEMM colonies formed from HSPCs edited for different targets (mean + SEM for n = 2). (C) Proportion of erythroid subsets day 20 of differentiation. The populations are gated as reticulocytes (CD235a + Hoechst-), erythroblasts (CD235a + Hoechst+), and pyrenocytes (CD235a-Hoechst+). (mean + SEM for n = 2). (D) Representative flow cytometry plots for HbF + ve cells analysis for different conditions. (E) Percentage of HbF + ve cells in different editing conditions (mean + SEM for n = 2). (F) Ratio of γ-globin chains to β-like globins observed in RP-HPLC chain analysis (mean + SEM for n = 2). (G) Representative variant HPLC chromatogram traces for hemoglobin tetramer variant analysis of day 20 erythroblasts. The areas under HbF and HbA are shaded red and gray, respectively. (H) Percentage of HbF tetramers detected by HPLC analysis (mean + SEM for n = 2)

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Fig. 8
figure 8

Graphical abstract of the work. The HSPCs were electroporated with Cas9-RNP and ssODN to introduce two naturally occurring beneficial mutations (-175T > C and − 158 C > T) and a PAM shield mutation (-153G > C). The electroporated HSPCs were then co-cultured with BM-MSCs and AZD-7648 small molecule. This process resulted in HBG promoter activation, with over 70% of HSPCs containing all three conversions, minimal indels, and no compromise on cell viability. The erythroblasts from the edited HSPCs had high HbF levels

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Discussion

Reactivating HbF by modifying the HBG promoter is a promising therapeutic strategy for treating β-hemoglobinopathies [45]. Cas9 RNP-mediated indels and base editing-mediated nucleotide changes successfully activate the HBG promoter, leading to the production of HbF tetramers [13, 14]. In this study, we demonstrate the simultaneous introduction of two productive mutations, -175T > C and − 158 C > T, into HSPCs using a ssODN-based approach, achieving up to 57% efficiency in vivo and resulting in functional HbF production.

We chose the ssODN-based template correction method because of its simplicity, scalability, and ability to incorporate multiple point mutations with a single set of reagents. ssODNs facilitate repair through the single-strand template repair (SSTR) pathway [35], ensuring precise modifications without unintended nucleotide changes or vector sequence integration issues [46]. Despite the previous AAV6-based HDR approach achieving less than 4% HDR at the HBG promoter in vivo, we optimized our method to overcome these limitations [16]. An asymmetric ssDNA design, longer toward the PAM proximal end and complementary to the target/gRNA binding strand, proved most effective for HDR conversion efficiency. This design’s success suggests a strand bias in ssDNA-mediated repair at the HBG1/HBG2 locus, a finding that requires validation with other loci in HSPCs. Intriguingly, we also found that 3’ end modification is sufficient to protect ssODNs. Genome-wide screening identified TREX1, an exonuclease with 3’-to-5’ hydrolytic activity, as a key determinant of HDR efficiency [47]. The 3’ end protection of our ssODN likely counteracts the nuclease activity of TREX-1, protecting the ssODN and leading to a longer period of retention within the electroporated cells.

Our small molecule screening revealed that AZD-7648 significantly improved the − 175T > C and − 158 C > T conversion efficiency, with a concomitant reduction in InDels. Concurrent with our studies, two independent screens also identified AZD-7648 as a candidate that strongly influences HDR outcomes [48, 49]. We further showed that AZD-7648 treatment does not affect the engraftment or multilineage potential of HSPCs (Fig. 4A-C). Our HSPC culture medium contained an RUS cocktail, which we previously reported for primitive stem cell maintenance [30] and including AZD-7648 in this regimen benefits HDR-mediated editing of primitive HSCs.

The − 175T > C mutation, a potent naturally occurring hereditary HPFH mutation, and the − 158 C > T polymorphism, known to induce HbF, were co-introduced. Technically,

The close proximity of -158 C > T to the Cas9 cut site resulted in a higher conversion rate than that of the − 175T > C modification, which is farther from the cut site. This led to up to 27% of HSPCs containing only the − 158 C > T mutation in vivo, in addition to HSPCs containing both mutations, which can potentially amplify the therapeutic effect. The combined approach utilizing complete HDR (HSPCs with both − 158 C > T & -175T > C modification), partial HDR (HSPCs with either − 158 C > T or -175T > C modification), optimized ssODN design, RUS cocktail, AZD-7648 treatment and BM-MSC coculture achieved HDR gene editing efficiency levels comparable to those previously attainable only through NHEJ-based gene editing methods.

Recently, the − 175 A > G modification at the HBG promoter was reported using AB8e, which achieved 60% base editing efficiency ex vivo but also resulted in significant bystander mutations (11% and 28%), with one potentially negating the beneficial effects of the − 175 A > G mutation. Approximately 42% of the − 175 A > G base-edited cells were retained in vivo, which reversed the sickling phenotype [14]. Our approach achieved up to 57% HDR-edited cells in vivo with no bystander mutations. However, we need to address variations in engraftment efficiency between animals and a 4.9 kb deletion in the homologous HBG2 promoter to achieve a base editing-equivalent approach for uniform HbF induction. Studies have shown that such 4.9 kb deletions are inevitable during HBG promoter editing with nucleases or with nickases [13, 50, 51]. The encouraging clinical data from ongoing RUBY and EdiTHAL trials, which employ AsCas12a nuclease for HBG promoter editing, support the use of the nuclease for HBG promoter editing [52, 53]. Deletions initiated at the − 115 site disrupt the BCL11A binding site and subsequently lead to HbF activation, which reportedly does not have deleterious effects on HSPCs [13, 50]. In our approach, the deletion initiates from the − 158 position and is not associated with any significant levels of HbF activation (Fig. 2B-E, RNP), and HbF activation is purely from HDR edits.

Our complete HDR efficiency reached over 80% with minimal indels in small-scale in vitro experiments. However, scaling up for mouse transplantation studies reduced the editing efficiency, indicating the need for further optimization of the cell dose, RNP, and ssODN ratio during the preparation of edited cell products in bulk. The sgRNA used in the study was recently reported to have no off-target effects according to guide seq [50]. However, the off-target effects associated with transient global suppression of NHEJ DNA repair by AZD-7648 have yet to be studied. Although our study indicated that AZD-7648 treatment has a negligible impact on the long-term repopulation of HSPCs (Fig. 4 and Supplementary Fig. 6A), any potential increase in mutation burden requires further investigation. Additionally, treatment with AZD-7648, which almost completely abolishes InDel formation, also increased the frequency of the 4.9 kb deletion on comparison with HDR edited cells that were not treated with AZD-7648 (Supplementary Fig. 4C). Whether it promotes much longer deletions needs to be investigated.

A significant limitation of our approach is the cell toxicity following ssODN editing. We mitigated this issue by coculturing HSPCs with bone marrow MSCs, which increased the absolute HDR cell frequency four-fold in vitro. Notably, BM-MSCs exhibited better results than WJ-MSCs, possibly because they resemble the component of anatomical niche of adult HSCs and promote better recovery from stress. Recently, it was shown that gene-edited HSPCs with Cas9 RNPs had increased expression of P21, a cell cycle regulator, and proinflammatory cytokines such as CXCL8, MCP-1, and TNF-α [46]. When used as a stromal coculture system for gene-edited HSPCs, MSCs reduce the activation of most of these factors and release pro-survival factors such as IL-10 and IL-18, improving transplantation efficiency [44]. We anticipate that MSC coculture may negate variable engraftment efficiency in vivo. However, the use of MSCs as stromal cells complicates the clinical manufacture of gene-edited cells. Alternative strategies to harness the beneficial effects of MSCs include chemical formulation of the culture media or the use of immortalized GMP-compliant engineered MSCs that secrete all the necessary factors for HSPCs.

Conclusions

Overall, our work serves as a proof of concept for employing ssODNs to introduce two beneficial mutations at a high frequency to treat β-hemoglobinopathies. This study paves the way for incorporating additional beneficial mutations and exploring the outcomes of all possible nucleotide conversions, which fall well within the ssODN sequence, to further enhance therapeutic efficiency.

With the necessity of preclinical safety studies, the promising results we present here have potential for clinical translation as therapeutics for both SCD and β-thalassemia. This could be a potential alternative to physically demanding treatment regimens and thus enhance quality of life by offering a lasting cure for patients with β-hemoglobinopathy.

Data availability

All the sequences corresponding to ssODNs, gRNAs, and oligos are provided in the supplementary materials section.The sequencing analysis files are available from Dryad data repository, which can be accessed by following the link: https://doiorg.publicaciones.saludcastillayleon.es/10.5061/dryad.nk98sf83j. Any other relevant materials used and/or analyzed during the current study are available from the corresponding author upon reasonable request.

Abbreviations

HSPCs:

Hematopoietic stem and progenitor cells

ssODN:

Single–stranded oligonucleotide

HDR:

Homology–directed repair

NHEJ:

Nonhomologous end joining

HbF:

Fetal hemoglobin

SCD:

Sickle cell disease

MSCs:

Mesenchymal stem cells

RNP:

Ribonucleoprotein

PAM:

Protospacer adjacent motif

HPLC:

High–performance liquid chromatography

HPFH:

Hereditary persistence of fetal hemoglobin

NBSGW:

NOD. Cg–KitW–41J Tyr+ Prkdcscid Il2rgtm1Wjl/ThomJ

DNA-PK:

DNA–dependent protein kinase

BM:

Bone marrow

WJ:

Wharton’s Jelly

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Acknowledgements

The authors acknowledge the funding support from the Department of Biotechnology, Ministry of Science and Technology, Government of India through the grants; BT/PR31616/MED/31/408/2019, BT/PR45683/MED/31/465/2022 and BT/PR26901/MED/31/377/2017 to ST. We thank CSIR-JRF fellowship for PB, ICMR-SRF fellowship for VV. The staff of the flow cytometry, animal facility, and core facilities of CSCR for their support. We thank Dr. R V Shaji and Dr.Poonkuzhali Balasubramanian for their expert advice and HPLC experiments and Dr. Rekha Pai for ddPCR analysis. We also thank Dr.Saranya Srinivasan, Dr.Karthik V Karuppusamy, Sumathi Rangaraj, Anant Kumar, and Anila George for their intellectual and technical input in this work. Use of AI: We acknowledge the use of chat GPT to improve language clarity.

Funding

The authors thank the funders of the Department of Biotechnology, Ministry of Science and Technology, Government of India (BT/PR31616/MED/31/408/2019, BT/PR45683/MED/31/465/2022 and BT/PR26901/MED/31/377/2017 to ST.

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Author notes

  1. Prathibha Babu Chandraprabha and Manoj Kumar K. Azhagiri contributed equally to this work and Shared first authorship.

Authors and Affiliations

  1. Centre for Stem Cell Research (CSCR), A Unit of InStem Bengaluru, Christian Medical College Campus, Vellore, Tamil Nadu, 632002, India

    Prathibha Babu Chandraprabha, Manoj Kumar K. Azhagiri, Vigneshwaran Venkatesan, Kirti Prasad, Sevanthy Suresh, Srujan Marepally, Alok Srivastava, Kumarasamypet Murugesan Mohankumar & Saravanabhavan Thangavel

  2. Manipal Academy of Higher Education, Manipal, Karnataka, 576104, India

    Prathibha Babu Chandraprabha, Manoj Kumar K. Azhagiri, Vigneshwaran Venkatesan, Kirti Prasad & Sevanthy Suresh

  3. Children’s Hospital Oakland Research Institute, UCSF Benioff Children’s Hospital Oakland, Oakland, CA, 94609, USA

    Wendy Magis & David I. K. Martin

  4. Department of Hematology, Christian Medical College, Vellore, Tamil Nadu, 632004, India

    Aswin Anand Pai & Alok Srivastava

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  1. Prathibha Babu Chandraprabha
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Contributions

Conceptualization: S.T., D.I.M., and W.M. Experimental execution and analysis: P.B., M.K.A., V.V., K.P., S.S., and A.A.P. Technical supervision: S.T., D.I.M., K.M.M. and S.K. M. Manuscript – review & editing: S.T.,D.I. M, P.B., M.K.A., V.V., K.M.M.,S.K. M and A.S. Funding acquisition: S.T.

Corresponding author

Correspondence to Saravanabhavan Thangavel.

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Ethical approval

The study titled “ In vivo efficacy and safety studies of CSCR-ST04, the gene edited autologous hematopoietic stem cells for the gene therapy of -hemoglobinopathies” was approved by the IRB of the Christian Medical College Vellore, with the approval number “IRB: 11807 (other) dated: 30.01.2019” and, the animal experiments were approved by the IAEC (approval number: 01/2019).

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Chandraprabha, P.B., Azhagiri, M.K.K., Venkatesan, V. et al. Enhanced fetal hemoglobin production via dual-beneficial mutation editing of the HBG promoter in hematopoietic stem and progenitor cells for β-hemoglobinopathies. Stem Cell Res Ther 15, 504 (2024). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s13287-024-04117-0

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Keywords

Stem Cell Research & Therapy

ISSN: 1757-6512