Introduction
Since the early 2000s, granulocyte colony-stimulating factor (G-CSF) has been routinely used to mobilize hematopoietic stem cells (HSCs) into the bloodstream for collection via leukapheresis to treat a wide variety of hematological as well as non-hematological diseases and disorders either in the autologous or allogeneic setting. 1 , 2 Essential for the success of this treatment strategy is the efficient mobilization of CD34 + hematopoietic stem and progenitor cells (HSPCs), reaching the widely accepted threshold of at least 2 × 10 6 but preferentially greater than 10 × 10 6 CD34 + HSPCs per kilogram of body weight. Although G-CSF mobilization has been successfully carried out millions of times over the last two decades, only about a third of all patients can reach this threshold with a single round of mobilization and leukapheresis, requiring a repeat collection. 3 To reduce the need for remobilization, G-CSF is often combined with the CXCR4 antagonist plerixafor (AMD3100), which boosts success rates for single leukapheresis transplants to almost two-thirds of patients. 4 , 5
However, approximately one-third of all patients respond poorly or remain unresponsive to G-CSF, 6 and even the additional use of plerixafor in this population often will not help to increase the CD34 yield. Furthermore, G-CSF has short-term side effects, including temporary bone and muscle pain, in more than 90% of the donors/patients and, less often, severe consequences, such as splenic rupture or pulmonary infiltrates. Especially problematic is that G-CSF can cause severe side effects in sickle cell disease [SCD] patients 7 , 8 , 9 or mobilization/collection of HSCs for transplant is difficult to achieve due to their low frequency of HSCs in pateients with Fanconi anemia and deficiency of adenosine deaminase 2. 10 , 11 , 12
To overcome these side effects and improve HSC mobilization, a great variety of compounds and small molecules have been developed and tested in the past years, each with their own unique method of action and often used in combination with previously established compounds. While some of these drugs have started to enter pre-clinical testing in healthy individuals, a more detailed assessment and side-by-side comparison with the traditional and well-established mobilization regimens are either limited to small animal models such as rodents or lacking entirely. Here, we comprehensively compared the mobilization of HSCs either using the gold standard G-CSF alone or in combination with plerixafor (AMD3100) and the CXCR2 ligand MGTA-145/GroβT (GroβT) 13 in the pre-clinical nonhuman primate (NHP) stem cell transplantation and gene therapy model. We assessed each combination’s mobilization efficiency, ability to leukapherese and enrich CD34 + HSPCs, as well as the capacities to gene-modify and transplant cells after myeloablative conditioning, closely mimicking the treatment of an SCD patient undergoing HSC gene therapy.
Results
AMD/GroβT efficiently mobilizes CD34 + CD90 + HSCs in rhesus macaques
To evaluate the mobilization efficiency of CD34 + HSPCs and CD34 + CD90 + HSCs, 4 animals each were treated with either 50 μg/kg G-CSF or 1 mg/kg AMD3100 and 450 μg/kg GroβT (AMD/GroβT) ( Figure 1 A; Table S1 ). G-CSF was administered for 5 days, while AMD3100 was given 1 h and GroβT 10 min prior to leukapheresis.
To initially determine the efficiency of mobilization, blood was analyzed by flow cytometry prior and during leukapheresis to assess the frequency of circulating CD34 + HSPCs and the HSC-enriched CD34 + CD90 + subset ( Figure 1 B). Both mobilization regimens demonstrated no significant differences in the frequency of CD34 + cells (G-CSF, 0.08% ± 0.06%; AMD/GroβT, 0.08% ± 0.05%) and the CD90 + subset (G-CSF, 0.05% ± 0.03%; AMD/GroβT, 0.04% ± 0.02%) ( Figure 1 C). Over 50% of all mobilized CD34 + cells co-expressed CD90, indicating a highly efficient and preferential release of primitive HSCs into the blood with either G-CSF alone or AMD/GroβT.
Animals were apheresed for up to 6 h with peripheral blood (PB) samples collected longitudinally for complete blood cell (CBC) counts, determine frequencies of CD34 + /CD90 + cells by flow cytometry, and calculate the absolute number of HSPCs and HSCs per microliter over time ( Figure 1 D). Applying clinical thresholds, successful mobilization of more than 5 CD34 + HSPCs per microliter of blood was achieved in 3 of 4 G-CSF animals, ranging from 5.1 (animal A18096) to 36.9 (A18133) CD34 + cells/μL ( Figure 1 D, top row, black lines). Throughout leukapheresis, the frequency of CD90 + HSCs within mobilized CD34 + HSPCs remained above 40% at all times, ranging from 2.1 (A19116) to 15.9 (A18096) HSCs/μL ( Figure 1 D, red lines). Of note, animal A18095 showed only a slight increase of CD34 + cells (maximum 2.4/μL) and CD90 + HSCs (maximum 0.66/μL) from baseline (CD34 + , 0.37/μL; CD90 + , 0.04/μL) and was therefore classified as a poor G-CSF responder and re-enrolled into mobilization with AMD/GroβT.
Very consistent mobilization was achieved in all 4 animals injected with AMD/GroβT, ranging from 7.2 (A18095) to 20.6 (A18097) CD34 + HSPCs/μL and 3.0 (A18095) to 9.0 (A18097) CD90 + HSCs/μL at peak mobilization ( Figure 1 D, bottom row). Kinetics of HSPC/HSC mobilization with AMD/GroβT were altered in comparison to G-CSF alone, with peak HSPC/HSC circulation seen 2–3 h post GroβT administration. Similar to G-CSF mobilization, the frequency of CD90 + HSCs within mobilized CD34 + HSPCs remained above 40% at all times. Most importantly, the poor G-CSF responder A18095 successfully mobilized CD34 + /CD90 + cells within the range of a G-CSF responder (>5 CD34 + /μL) upon AMD/GroβT treatment.
These results highlight both fast and consistent mobilization of CD34 + HSPCs and the HSC-enriched CD34 + CD90 + subset with AMD/GroβT without any significant differences compared to G-CSF alone. Most importantly, AMD/GroβT allowed the remobilization of a poor G-CSF responder.
Comparable CD34 + and CD90 + HSCs counts per kilogram of body weight after G-CSF and AMD/GroβT mobilization
To allow ex vivo gene editing of HBG , CD34 + cells were enriched from the leukapheresis products after G-CSF or AMD/GroβT mobilization as described previously. 14 CD34 products were comprehensively analyzed for their phenotypic composition using flow cytometry.
Comparable purities of CD34 + HSPCs were reached for leukapheresis-derived CD34 products with both mobilization regimens ( Figures 2 A and 2B). As observed previously in the PB samples ( Figure 1 B), CD90 + HSCs were highly enriched in CD34 products after leukapheresis (G-CSF, 32 ± 7.09%; AMD/GroβT, 38.88 ± 9.95%) ( Figures 2 A and 2B). Quantification of CD34 + HSPCs based on the animal’s weight showed similar cell yields from G-CSF- and AMD/GroβT-mobilized leukapheresis products (G-CSF, 2.4 ± 1.6 × 10 6 ; AMD/GroβT, 2.2 ± 1.2 × 10 6 ; Figure 2 C). Equally comparable was the absolute number of CD90 + HSCs per kilogram of body weight (G-CSF, 8.4 ± 0.69 × 10 5 ; AMD/GroβT, 7.8 ± 2.1 × 10 5 ; Figure 2 C).
To confirm that the phenotyping of CD34 subsets with CD90 and CD45RA described previously in steady-state bone marrow (BM) and G-CSF/stem cell factor (SCF)-primed BM similarly applies to G-CSF- and AMD/GroβT-derived HSPC products, CD34 subsets were purified by fluorescence-activated cell sorting (FACS) and introduced into colony-forming cell (CFC) assays to assess their clonogenic erythro-myeloid differentiation potential. As described previously, erythroid differentiation potential was almost exclusively limited to CD34 + CD90 – CD45RA – erythro-myeloid progenitors (EMPs), CD34 + CD90 – CD45RA + lympho-myeloid progenitors (LMPs) were restricted to myeloid differentiation, and the HSC-enriched CD34 + CD90 + CD45RA – population was highly enriched for granulocytic cell fates mixed with cobblestone-forming colonies ( Figure 2 D). Furthermore, overall colony formation rates for G-CSF- and AMD/GroβT-derived CD34 subsets were closely matching previously described frequencies seen in steady-state BM and G-CSF/SCF-primed BM, ranging from 10% to 25% depending on the CD34 subset.
In summary, CD34 products from G-CSF- and AMD/GroβT-derived leukapheresis products yield comparable CD34 + cell counts per kilogram of body weight and are highly enriched for CD90 + HSCs.
Efficient ex vivo gene modification of AMD/GroβT-mobilized CD34 + HSPCs
Isolated G-CSF- and AMD/GroβT-mobilized CD34 + cells were cultured overnight, followed by electroporation with CRISPR-Cas9 mRNA and HBG -specific guide RNA (gRNA) to reactivate fetal hemoglobin as described previously. 15 After overnight culture, gene-modified cells were comprehensively quality controlled regarding their phenotypic composition, maintenance of their erythro-myeloid differentiation potential in CFC assays, and HBG editing efficiency.
Flow cytometry analysis of the gene-modified CD34 cell fraction showed a culture-induced loss of CD34 – cells and consequent increase in CD34 purity (G-CSF, 76.53% ± 11.86%; AMD/GroβT, 63.05% ± 3.39%) ( Figures 3 A and 3B). No changes were seen in the frequency of CD90 + HSCs within the CD34 fractions upon culture and editing (G-CSF, 37.3% ± 1.71%; AMD/GroβT, 36.5% ± 10.17%) ( Figures 3 A and 3B). Absolute CD34 + cell counts per kilogram of body weight declined for G-CSF and AMD/GroβT-mobilized products by about 30%–50% upon culture and gene modification (G-CSF, 1.7 ± 0.6 × 10 6 [−30%]; AMD/GroβT, 1.1 ± 0.5 × 10 6 [−50%]; Figure 3 C). CD90 + HSCs were lost in the process at a similar rate, with all collection methods showing, on average, very comparable yields (G-CSF, 6.4 ± 2.1 × 10 5 [−27%]; AMD/GroβT, 4.1 ± 2.4 × 10 5 [−52%]; Figure 3 C).
Next, gene-modified CD34 subsets were FACS purified and introduced into CFC assays to assess their total colony-forming and erythro-myeloid differentiation potential. No noticeable differences were seen in the overall colony-forming frequency (10%–25%) or in the cell-type-specific differentiation potential for the phenotypic CD34 subsets in comparison to freshly isolated CD34 + cells ( Figure 3 D).
Finally, bulk CD34 + cells were analyzed for the editing efficiency of the HBG locus using Sanger sequencing and Tracking of Indels by Decomposition (TIDE) analysis ( Figure 3 E). On average, over 70% editing was seen for G-CSF- and AMD/GroβT-mobilized (G-CSF, 76% ± 6.7%; AMD/GroβT, 66% ± 14%; Figure 3 E). DNA from one G-CSF- and three AMD/GroβT infusion products were further subjected to next-generation sequencing (NGS) to characterize the pattern of insertions or deletions (indels) at the HBG promoter ( Figure 3 F). As seen previously, a diverse repertoire of deletions ranging from 1 to 13 nt was seen in CD34 + HSPCs, including the naturally occurring, highly potent 13-nt deletion seen in the benign genetic condition known as hereditary persistence of fetal hemoglobin (HPFH). 16
These data confirm that G-CSF- and AMD/GroβT-derived CD34 + cells can be efficiently isolated, cultured, and CRISPR-Cas9 edited ex vivo without compromising the phenotypic composition or erythro-myeloid differentiation potential. All generated cell products of gene-modified CD34 + HSPCs qualified for the transplantation into myeloablative conditioned animals.
Rapid transfusion-independent multilineage recovery after transplantation with both mobilization regimens
Upon successful quality control of the infusion products, NHPs were myeloablation conditioned, receiving fractionated total body irradiation (1,020 cGy) ( Figure 1 A). Of note, one of the three G-CSF animals was removed from the study after leukapheresis due to unrelated health issues. The follow-up for animal A19116 (G-CSF group) was terminated after 90 days due to infection complications.
After cell infusion, all animals were closely monitored for hematopoietic recovery, applying clinical parameters for neutrophil recovery (absolute neutrophil count > 0.5 K/μL) and platelet recovery (platelets > 20 K/μL). Recovery of neutrophils, platelets, and the need for platelet transfusions was compared side by side with our historic controls.
Animals receiving G-CSF-mobilized CD34 + cells recovered neutrophils after 9 ± 1.41 days and platelets after 13.5 ± 3.54 days ( Figures 4 A, 4B, and S1 ). In contrast, neutrophil recovery was slightly slower with AMD/GroβT at 12 ± 1.63 days, followed by platelets after 15.25 ± 1.89 days ( Figures 4 A, 4B, and S1 ). However, these differences were not significant. G-CSF- and AMD/GroβT-treated animals required 1.5 ± 0.71 and 1 ± 0.82, respectively, with one AMD/GroβT animal being entirely transfusion independent ( Figure 4 C).
CBC counts were monitored for all animals to ensure full recovery of all hematopoietic compartments back to the normal range. A rapid drop in white blood cells (WBCs) and platelets was seen shortly after conditioning, with most values being out of the normal range for up to 2 months post transplantation ( Figure 4 D). While neutrophils and platelets returned to the normal range within approximately 20–30 days for all animals, lymphocyte recovery in some animals demonstrated a variable delay of up to 3 months to full recovery ( Figure 4 D).
To obtain more detailed information on the multilineage reconstitution, including lymphocyte subsets, blood was analyzed by flow cytometry ( Figure 4 E). Granulocytes (CD11b + CD14 – SSC high ) and monocytes (CD11b + CD14 + SSC low ) showed a rapid recovery after transplantation, followed by T cells (CD3 + ), B cells (CD20 + ), and natural killer cells (CD16 + ) recovering within 4 months post transplantation ( Figure 4 E). A more detailed assessment of CD3 + T cells showed an even contribution of CD4 + cytotoxic and CD8 + regulatory T cells in the blood of all animals.
G-CSF- and AMD/GroβT-derived CD34 + cells demonstrated rapid engraftment and reconstitution potential after myeloablative hematopoietic stem cell transplant despite significantly smaller total CD34 doses per kilogram in comparison to our previously published work. 17 Recovery kinetics for neutrophils and platelets matched the previously described correlation of CD90 cell doses per kilogram of body weight seen for animals transplanted with primed BM CD34 + cells, 17 confirming that the CD90 subset enriched in G-CSF- and AMD/GroβT-mobilized CD34 + HSPCs were exclusively responsible for hematopoietic recovery and were of identical quality as seen previously from steady-state BM or other HSC sources.
Contribution of gene-modified HSCs to all peripheral blood lineages
Next, we investigated the engraftment of gene-modified HSCs and their contribution to the PB. Gene modification efficiency was initially determined in the PB by measuring the frequency of cells expressing fetal hemoglobin (F cells). Red blood cells (RBCs) were fixed, permeabilized, stained, and the percentage of fetal hemoglobin expressing (HbF) + RBCs measured by flow cytometry over time ( Figure 5 A). As seen in our previous studies, F cell frequencies rapidly increased within the first 2 months as a result of the conditioning, stress hematopoiesis, and contribution of gene-modified HSPCs with a peak level at 36.55% ± 9.12% and 27.97% ± 6.03% for G-CSF and AMD/GroβT, respectively. Due to the previously described non-lethal conditioning and competition with surviving endogenous, unedited HSCs, the F cell frequency gradually declined, and F cell frequencies started stabilizing after 4–5 months at 2.02% ± 1.16% ( n = 3) in AMD/GroβT animals and 21.65% ± 9.26% ( n = 1) for the remaining G-CSF animal.
To verify that occurrence of F cells was due to the gene editing of HSCs and not a stress-related response leading to a transient production of F cells, gene editing frequencies in PB WBCs were determined at the genomic level using NGS ( Figure 5 B). Similar to the peak F cells frequency in the PB, greatest HBG editing in PB WBCs was seen within the first 2 months post transplantation.
Peak editing of 76.54% ± 6.82% and 68.37% ± 15.43% for G-CSF and AMD/GroβT, respectively, was recorded immediately after cell infusion, matching the previously determined editing efficiency in the infusion products ( Figure 3 D). The frequency of HBG -edited PB WBCs stabilized after 2 months at 7.33% ± 3.03% ( n = 3) in AMD/GroβT animals and 18.94% ± 0.45% ( n = 1) for the remaining G-CSF animal. More detailed analysis of the CRISPR-Cas9-mediated indels at the HBG locus revealed nearly identical patterns for G-CSF and AMD/GroβT animals, with the pattern of indels remaining very consistent long term ( Figure 5 C). Of note, the frequency of the 13-nt deletion described to induce the highest level of HbF expression was similarly represented in both cohorts at about 10% of all indels and persisted throughout the entire follow-up. These data confirm successful long-term engraftment of G-CSF- as well as AMD/GroβT-mobilized and ex vivo gene-edited HSCs after myeloablative conditioning.
Full recovery and stable long-term engraftment of gene-modified HSCs in the bone marrow stem cell compartment
Long-lasting production of F cells requires efficient engraftment of gene-modified HSCs in the BM stem cell compartment. To confirm homing and persistence of gene-modified HSCs in the BM of transplanted animals, BM samples were collected 6 and 12 months post transplantation. BM specimens were initially analyzed for their phenotypical composition and frequency of HSPCs as well as HSCs in comparison to our historic controls ( Figures 6 A and 6B). No significant differences were seen in the phenotypic HSPC composition ( Figure 6 A) or the overall frequency of CD34 + HSPCs and CD90 + HSCs ( Figure 6 B) when comparing the G-CSF and AMD/GroβT animals among each other. CD34 + frequencies in G-CSF animals reached 0.63% after 6 months and increased to 0.87% after 12 months. CD34 + frequencies were slightly higher at 6 months in AMD/GroβT animals at 1.06% ± 0.66%, with no further increase after 12 months, showing 1.05% ± 0.54%. Less variability was seen for CD90 + HSC frequencies averaging 0.084% and 0.09% ± 0.05% for G-CSF and AMD/GroβT, respectively. CD90 frequencies continued to increase in both groups to 0.13% and 0.1% ± 0.02% for G-CSF and AMD/GroβT, respectively.
To verify that phenotypic HSPCs and HSCs retain their erythro-myeloid differentiation potential, CD34 + HSPCs and CD90 + HSCs were FACS purified for CFC assays. Robust multilineage differentiation potential was seen for both CD34 + HSPCs (G-CSF, 21.3% ± 2.2%; AMD/GroβT, 11.1% ± 5.6%) as well as CD90 + HSCs (G-CSF, 20.2% ± 0.4%; AMD/GroβT, 16.6% ± 3.9%; Figure 6 C), closely matching data from our historic control animals. As expected, bulk CD34 + HSPCs showed erythroid and myeloid differentiation biased toward mixed myeloid colonies due to the strong representation of CD34 + CD90 – CD45RA + cells ( Figure 6 A), whereas CD90 + HSCs primarily generated granulocytic colonies. Overall, CFC potential in bulk CD34 + HSPCs (AMD/GroβT, 17.5% ± 5.4%) and CD90 + HSCs (AMD/GroβT, 25.2% ± 8.5%) remained stable in AMD/GroβT animals at 12 months.
Finally, FACS-purified BM lineages (monocytes, granulocytes, T cells, B cells, natural killer cells, and erythroid progenitors), CD34 + HSPCs, and CD34 subsets, including CD90 HSCs, were analyzed by NGS to confirm persistence of gene-modified HSCs and even contribution of HSCs to all progenitors and lineages ( Figures 7 A and 7B). HBG editing was seen across all FACS-purified lineages as well as HSPC subsets in both G-CSF and AMD/GroβT animals ( Figure 7 C). Total editing efficiency in the FACS-purified lineages and HSPC subsets matched the previously observed editing efficiency seen long term in bulk PB WBCs, which was approximately 20% for the G-CSF-mobilized animal A18131 and 5%–10% for the AMD/GroβT animals A17283 and A18095 with the longest follow-up available ( Figure 5 B). Patterns of indels were similarly diverse as seen previously in PB WBCs, and the 13-nt HPFH deletion persisted in all samples, including the FACS-purified CD34 + CD90 + subset, demonstrating no significant difference when comparing G-CSF and AMD/GroβT animals, indicating stable long-term engraftment of gene-edited HSCs.
Discussion
In this study, we demonstrated rapid and efficient mobilization of HSCs using AMD3100 in combination with GroβT for ex vivo gene therapy. In comparison to G-CSF mobilization, less variable mobilization of CD34 + HSPCs and CD90 + HSCs was observed with AMD/GroβT, including the ability to efficiently mobilize a poor G-CSF responder. No qualitative or quantitative differences were seen for the gene-modified infusion products or in the short-term recovery and long-term multilineage engraftment potential of G-CSF- and AMD/GroβT-mobilized HSCs when comparing the two regimens.
Traditional HSC mobilization with G-CSF has long been successful, enabling millions of allogeneic and autologous transplants. 18 However, the known side effects of G-CSF, the high rate of non-mobilizers, and the need for G-CSF-free mobilization regimens for patients suffering from SCD sparked the search for alternative mobilization many years ago. A wide range of drugs to either block CXCR4/CXCL12 adhesion (e.g., balixifortide, Me6, and motixafortide), inhibit integrin function (e.g., BIO5192, firategrast, and N-(benzenesulfonyl)-L-prolyl-L-O-(1-pyrrolidinylcarbon)tyrosine [BOP]), and others (FG4592, sildenafil citrate, and desipramine) have been tested in small and large animal models for their ability to mobilize HSCs into the periphery. 19 , 20 , 21 , 22 , 23 , 24 Most of these drugs were primarily tested as an add-on for G-CSF to either improve mobilization above the needed threshold or reduce the rate of G-CSF non-responders. G-CSF-free mobilization using newer drugs, either alone or in combination with other drugs, is tested less often.
However, with the development of gene therapy strategies for SCD, the search for G-CSF-free mobilization regimens has gained more traction, as G-CSF can cause a sickling crisis due to the induced leukocytosis. To avoid these complications, mobilization exclusively with AMD3100 has been successfully tested in clinical trials for efficiency, safety, and feasibility to treat SCD patients and, more recently, become one of the first US Food and Drug Administration-approved HSC gene therapy strategies. 25 , 26
Although successful, AMD3100 alone has shown lower rates of HSPC mobilization across multiple experimental and pre-clinical studies. 27 , 28 , 29 , 30 As a benchmark, 20 CD34 + cells per microliter of blood is considered the ideal goal to achieve at least 5 × 10 6 CD34 + cells per kilogram of body weight and avoid multiple mobilizations. 31 To accomplish this rate at peak mobilization, at least 80 μg/kg AMD3100 needs to be administered, ideally 160 μg/kg or even 240–320 μg/kg, to allow continuous collection of CD34 + cells throughout the leukapheresis 32 , 33 , 34 ; however, even at this higher dose, many patients still require two leukapheresis collections to yield sufficient CD34 + cells for transplantation. 28 , 29 , 30 To boost AMD3100-only mobilization (without the use of G-CSF) and overcome issues with potential AMD3100 non-responders, 7 administration of additional compounds together with AMD3100 has been investigated. 19 The most successful and fastest strategy so far has been the combination of AMD3100 with GroβT. 35 Markedly increased mobilization rates of CD34 + cells were seen when using GroβT in combination with G-CSF 36 or AMD3100 35 that exceeded the previously observed frequencies with the combination of G-CSF and AMD3100. Following these pre-clinical studies, mobilization of HSCs in multiple myeloma patients was successfully carried out using AMD and GroβT. 37 Here, we show that AMD3100/GroβT-mobilized HSCs can be harvested by leukapheresis in NHPs for gene therapy and transplantation, further expanding the potential use of this mobilization regimen.
Crucial for the success of HSC gene therapy is the successful mobilization and collection of phenotypically primitive HSCs for ex vivo modification. In prior studies, we established that the purification and gene modification of the HSC-enriched CD34 + CD45RA – CD90 + cell fraction in humans and NHPs directly correlates with the onset of recovery and is solely responsible for short-as well as long-term hematopoietic reconstitution in the autologous, 15 , 17 , 38 allogeneic, 39 and xenograft setting. 40 , 41 Here, we comprehensively assessed the cell numbers, phenotypical composition, and quality of cell products throughout the entire process of mobilization/priming, leukapheresis/harvest, ex vivo gene modification, and infusion product preparation, comparing our historic control data from G-CSF/SCF-primed BM-derived cell products 42 with leukapheresis-collected cells after either G-CSF-only or AMD/GroβT mobilization. Although we collected significantly fewer CD34 + HSPCs when using mobilization and leukapheresis in comparison to harvesting primed BM, the overall quality of the CD34 + cell products was markedly improved and contained predominantly phenotypically primitive CD34 + CD45RA – CD90 + HSCs in the rhesus macaque. Throughout the process of ex vivo culture and gene editing, a similar loss of cells was observed across all three collection strategies without any obvious changes in the phenotypic composition. Consequently, animals mobilized with G-CSF only or AMD/GroβT did not receive the clinically recommended dose of more than 10 × 10 6 CD34 + HSPCs/kg, whereas historic G-CSF/SCF-primed animals, on average, were transplanted with 12 × 10 6 CD34 + HSPCs/kg. Regardless of this difference and as established previously to correlate with the onset of neutrophil and platelet recovery, 17 the average number of CD34 + CD45RA – CD90 + HSCs/kg was almost identical across all three groups, leading to neutrophil and platelet recovery within the same time range for all animals, as seen previously for our controls. Of note, AMD/GroβT mobilization led to a temporary spike in platelets post leukapheresis, shortening the duration of thrombocytopenia as well as significantly reducing the need for transfusions. Finally, as seen in patients, one of four G-CSF-mobilized animals did not respond but successfully mobilized HSPCs upon AMD/GroβT administration, providing an avenue for poor or non-responders to G-CSF.
A major focus of this study was to evaluate the ability to isolate, gene modify, and engraft mobilized CD34 + HSPCs. Although we achieved 60%–80% editing of the HBG -175 locus in G-CSF- as well as AMD/GroβT-mobilized CD34 + HSPCs, lower long-term multilineage engraftment of gene-modified lineages was seen with AMD/GroβT-mobilized cell products, whereas G-CSF-mobilized cells were within the same range as our historic controls. As reported previously in the mouse, the phenotype of mobilized HSCs was more primitive and showed increased multilineage engraftment potential, 36 suggesting that AMD/GroβT-mobilized HSCs in the NHP are similarly immature and/or quiescent, likely reducing the ability of gene editing, as the required DNA repair machinery is mostly inactive. 43 Extended priming of cells or cytokine treatment to initiate cell division and activate essential DNA repair proteins may be needed to increase gene editing levels in AMD/GroβT-mobilized HSCs. However, stimulation of cell proliferation may lead to loss of engraftment potential and differentiation of HSCs, requiring careful optimization of ex vivo gene editing protocols to find the balance between efficiency and HSC loss.
Assessment of the DNA repair mechanism comparing G-CSF and AMD/GroβT-mobilized HSCs may further shed light on the observed differences in engraftment of gene-modified cells. As described previously, due to the homology between HBG1 and HBG2 , a non-homologous end-joining (NHEJ)-mediated 4.9-kb deletion can occur that is negatively selected upon transplantation in the NHP. 42 As DNA repair in primitive HSCs is more likely to be NHEJ mediated, 44 , 45 lower engraftment in the AMD/GroβT group may be associated with higher rates of large deletions and, therefore, loss of edited HSCs. Similarly, assessment of the 13-bp deletion, a predicted microhomology-mediated end joining (MMEJ) product, could shed light on this, as MMEJ is less prevalent in more primitive HSCs and enriched in progenitors. 44 , 45 Although no significant difference in the frequency of engrafted CD90 + HSCs carrying the 13-bp deletion was seen when comparing G-CSF and AMD/GroβT animals, more detailed assessment of this editing pattern in the CD90 HSC subset of infusion products may help interpret the differences in outcome.
An avenue to entirely avoid the risk of HSC loss and bypass the extensive infrastructure for ex vivo HSC gene therapy has recently been tested in human CD46 transgenic mice mobilized with AMD3100 and GroβT (MGTA-145) using a helper-dependent adenovirus HDAd5/35 ++ vector system to correct thalassemia. 46 Similarly, SCD mice mobilized with the combination of AMD3100 and WU-106, an inhibitor of integrin α4β1, were successfully gene modified in vivo using adenoviral vectors carrying a prime editor. 47 A similar avenue for in vivo HSC gene therapy in mice, NHPs, as well as humans can be envisioned for the use of either HSC-targeted lentiviral vectors, 48 lipid nanoparticles, 49 or engineered virus-like particles 50 , 51 , 52 to deliver a broad variety of transgenes, mRNA therapeutics, or proteins/nucleases. Although some of these delivery vehicles may not require mobilization of HSCs for successful engineering, mobilization-mediated stimulation of HSCs may still be required to unleash the full potential of the delivered cargo.
In summary, the combination of AMD3100 with GroβT in the NHP large animal model did successfully mobilize a very primitive HSC population for ex vivo gene therapy and transplantation. The mobilization efficiency, cell yields, and reliability of this mobilization regimen allowed for successful transplantation of gene-modified HSCs in multiple animals without any obvious impact on the recovery or long-term multilineage engraftment. In particular, the rapid response will help significantly improve clinical pipelines, reduce the number of G-CSF non-responders, and enable safe mobilization of SCD patients for gene therapy. Follow-up studies will be needed to optimize the ex vivo editing of this more primitive and quiescent HSC subset as well as evaluate the opportunity to gene modify the cells directly in the blood stream for in vivo gene therapy.
Materials and methods
NHP studies
Healthy juvenile rhesus macaques were housed at the University of Washington (UW) National Primate Research Center (WaNPRC) under conditions approved by the American Association for the Accreditation of Laboratory Animal Care. All experimental procedures performed were reviewed and approved by the institutional animal care and use committee of the Fred Hutch Cancer Center and UW, protocol #3235-01. This study was carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health (the Guide ), including at least twice-daily observation by animal technicians for basic husbandry parameters (for example, food intake, activity, stool consistency, and overall appearance), as well as daily observation by a veterinary technician and/or veterinarian. Animals were housed in cages approved by the Guide and in accordance with Animal Welfare Act regulations. Animals were fed twice daily and were fasted for up to 14 h before sedation. Environmental enrichment included grouping in compound, large activity items, or run through connected cages, perches, toys, food treats, and foraging activities. If a clinical abnormality was noted by WaNPRC personnel, standard WaNPRC procedures were followed to notify the veterinary staff for evaluation and determination for admission as a clinical case. Animals were sedated by administration of ketamine HCl and/or Telazol and supportive agents for balanced anesthesia (such as diazepam and midazolam) before all procedures. After sedation, animals were monitored according to WaNPRC standard protocols. WaNPRC surgical support staff are trained and experienced in the administration of anesthetics and have monitoring equipment available to assist with electronic monitoring of heart rate, respiration, and blood oxygenation; audible alarms and digital readouts; and monitoring of blood pressure, temperature, etc. For minor procedures, the presence or absence of deep pain was tested by the toe pinch reflex, and the absence of response (leg flexion) to this test indicated adequate anesthesia. In cases of general anesthesia, similar monitoring parameters were used, and anesthesia was tested by the loss of palpebral reflexes (eye blink). Analgesics (generally buprenorphine with meloxicam or buprenorphine slow release) were provided as prescribed by the clinical veterinary staff for at least 48 h after the procedures and could be extended at the discretion of the clinical veterinarian based on clinical signs.
CD34 + enrichment and culture
For experiments utilizing mobilized peripheral blood and apheresis products, mobilization drugs were administered as outlined in Figure 1 A. Regular blood draws were performed before and during leukapheresis, lysed twice in ammonium chloride lysis buffer, and analyzed by flow cytometry.
Before enrichment of CD34 + cells, red cells were lysed in ammonium chloride lysis buffer, and WBCs were incubated for 20 min with the 12.8 immunoglobulin M anti-CD34 antibody, and then washed and incubated for another 20 min with magnetic-activated cell sorting anti-immunoglobulin M microbeads (Miltenyi Biotech). The cell suspension was run through magnetic columns enriching for CD34 + cell fractions with a purity of 60%–80%, confirmed by flow cytometry. Human CD34 + cells were harvested and enriched from mobilized PB as described previously. 53 Enriched CD34 + cells were cultured in SFEMII media (STEMCELL Technologies) supplemented with 100 U/mL penicillin and streptomycin (Gibco by Life Technologies) and SCF (PeproTech), thrombopoietin (PeproTech), and Fms-related tyrosine kinase 3 ligand (Miltenyi Biotec) (100 ng/mL for each cytokine).
HBG editing
Cas9 purified protein was obtained from Thermo Fisher Scientific (TrueCut Cas9 protein v2, 5 μg/μL, A36499), and chemically modified gRNAs (2′-O-methyl analogs and 3′ phosphorothioate internucleotide linkages at the first three 5′- and 3′-terminal RNA residues) were custom ordered from Synthego. Lyophilized gRNAs were resuspended in nuclease-free water at a concentration of 200 pmol/μL and stored as frozen aliquots at −80°C. For the editing of rhesus CD34 + or CD90 + subsets, cells were first cultured overnight after enrichment and/or sorting. CRISPR-Cas9 RNPs were formed by combining 180 pmol Cas9 protein with 1,800 pmol gRNA at room temperature for 10 min and used for the electroporation of 3 million cells in 2 mm cuvettes following the manufacturer’s instructions (Harvard Apparatus). Edited cells were recovered in medium overnight at 37°C before infusion into the animals.
Conditioning and recovery support
Rhesus macaques were conditioned with myeloablative total body irradiation of 1,020 cGy from a 6 MV X-ray beam of a single-source linear accelerator located at the Fred Hutch South Lake Union Facility (Seattle, WA, USA). The dose was administered at a rate of 7 cGy/min, delivered as a midline tissue dose. After transplant, G-CSF was administered daily from the day of cell infusion until the animals began to show onset of neutrophil recovery. Supportive care, including antibiotics, electrolytes, fluids, and transfusions, was given as necessary, and blood counts were analyzed daily to monitor hematopoietic recovery.
CBC counts
Post-conditioning numbers of total WBCs, platelets, neutrophils, and lymphocytes were determined by CBC count integrated with flow cytometry-based quantification of HSPC markers as described above. Separately, cell counts for CD34 + enrichment and infusion products were determined using the Countess II automated cell counter (Thermo Fisher Scientific) with trypan blue staining. CD34 + cell purity was confirmed by flow cytometry.
Flow cytometry and FACS
Antibodies used for FACS analysis and sorting of NHP cells are listed in Table S2 . Dead cells and debris were excluded by forward/side scatter (FSC/SSC) gating. Flow cytometry analysis was performed on an LSR IIu, FACSAria IIu, and Canto II (BD Biosciences). Cells for in vitro assays were sort-purified right after CD34 + enrichment using a FACSAria IIu cell sorter (BD Biosciences), and purity was assessed by recovery of sort-purified cells. Dead cells and debris were excluded by FSC/SSC gating.
HbF measurements
HbF was measured by flow cytometry after fixation and permeabilization of unlysed blood using procedures described previously. 54
CFC assays
A total of 600 FACS-purified cells were seeded into 3.5 mL ColonyGEL 1402 (ReachBio, Seattle, WA, USA) in 35-mm Petri dishes. Hematopoietic colonies were scored after 10–14 days. Arising colonies were identified as colony-forming unit (CFU) granulocyte (CFU-G), CFU macrophage (CFU-M), CFU granulocyte-macrophage (CFU-GM), and burst-forming unit-erythrocyte (BFU-E). Colonies consisting of erythroid and myeloid cells were scored as CFU-GEMM.
Quantifying indels with MiSeq
Editing at the HBG locus was measured either by TIDE analysis using Sanger sequencing 55 or by NGS using Illumina barcoded, 2 × 150 bp paired-end MiSeq primers for complete sequencing (Illumina). Primers are described in Table S3 , and bioinformatic processing of sequencing data was conducted using an in-house-built pipeline. 54
Statistics
Statistical analyses were performed in GraphPad Prism 10. Statistical tests were used for the data analysis as indicated in the figure legends.
Data availability
NGS data can be found under BioProject ID PRJNA1310223 on the NCBI Sequence Read Archive (SRA). All other data presented in this manuscript are available upon request.
Acknowledgments
Magenta Therapeutics provided GroβT and funding to carry out the study. We thank Helen Crawford for help in preparing this manuscript and figures. This work was supported in part by National Institutes of Health grants R01 HL136135 and R01 HL147324 (to H.-P.K.) and T32 HL007093 (to A.R.), Cancer Center Support Grant P30 CA015704, RRID:SCR_022613 (Flow Cytometry Shared Resource), of the Fred Hutchinson Cancer Center/University of Washington/Seattle Children’s Cancer Consortium, and P51OD010425 from the Washington National Primate Research Center. In addition, H.-P.K. is a Markey Molecular Medicine Investigator and holds the José Carreras/E. Donnall Thomas Endowed Chair for Cancer Research as well as the Stephanus Family Endowed Chair for Cell and Gene Therapy.
Author contributions
Conceptualization, S.R. and H.-P.K.; methodology, S.R., D.P., and M.R.E.; visualization, A.R. and S.R.; funding acquisition, H.-P.K.; project administration, S.R. and H.-P.K.; supervision, S.R. and H.-P.K.; writing – original draft, S.R., A.R., and H.-P.K.; writing – review & editing, all authors.
Declaration of interests
S.R. is a consultant to Inograft Inc. and Ensoma Inc. H.-P.K is or was a consultant to and has or had ownership interests with Rocket Pharmaceuticals, Homology Medicines, VOR Biopharma, and Ensoma Inc. H.-P.K. was also a consultant to CSL Behring and Magenta Therapeutics.
Footnotes
Supplemental information can be found online at https://doi.org/10.1016/j.omtm.2025.101558 .
Contributor Information
Hans-Peter Kiem, Email: hkiem@fredhutch.org.
Stefan Radtke, Email: sradtke@fredhutch.org.
Supplemental information
Document S1. Figures S1 and Tables S1–S3
Document S2. Article plus supplemental information
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Associated Data
Supplementary Materials
Document S1. Figures S1 and Tables S1–S3
Document S2. Article plus supplemental information
Data Availability Statement
NGS data can be found under BioProject ID PRJNA1310223 on the NCBI Sequence Read Archive (SRA). All other data presented in this manuscript are available upon request.