Application Note
Advancing a CCR5 Gene Editing Therapy to Clinical Trial: Overcoming Viral Vector Toxicity
Background
HIV infection is currently managed by lifelong antiretroviral therapy (ART), a modality that is associated with chronic toxicity, challenging patient compliance, and comes at a significant cost over a patient’s lifetime. Creating an HIV-1 infection-resistant immune system via gene editing has shown preclinical success and offers hope for a single dose therapeutic alternative.1, 2
Both preclinical and clinical studies have demonstrated that blocking or mutating CCR5, an R5-tropic HIV-1 coreceptor — whether through small molecule inhibition, presence of a natural mutation, or therapeutic gene modification — can render cells resistant to HIV-1 infection.3- 7 In one instance, a patient was "cured" following transplant of allogeneic stem cells containing a bi-allelic CCR5 mutation (CCR5Δ32/Δ32) with no evidence of HIV for eight years despite the halting of ART.8 , 9 Allogeneic stem cell transplant as a routine HIV therapy is limited by the availability of HLA-matched CCR5Δ32/Δ32 donors and comes with a high risk of morbidity and mortality.10
Adoptive transfer of autologous CD4+ T cells following ZFN-mediated CCR5 disruption (SB-728-T) was shown to be clinically safe, with cells engrafting and persisting over time.2 The limitation of this approach, however, is that patient monocytes — a population of immune cells believed to be a key reservoir for HIV infection — maintain wild-type CCR5. In contract, CCR5 gene editing of autologous HPSCs has the potential to result in HIV-1-resistant immune cells of multiple lineages, including both CD4+ T cells and monocytes, throughout the patient’s lifespan.
Disruption of CCR5 in adult mobilized CD34+ cells using ZFN in a clinical setting has been reported. Unfortunately, the cytotoxicity of the adenoviral vector used to deliver the ZFN machinery prevented its use in the intended clinical trial.5 It was determined that to move the clinical trial forward a non-viral, clinically-feasible, and regulatory-compliant technology was necessary to deliver the ZFN gene editing machinery to HSPCs.
Aim
Conduct preclinical IND-enabling studies for the clinical-scale manufacturing of CCR5-disrupted autologous HSPCs via non-viral delivery of SB-728 mRNA using the MaxCyte GT in support of an anticipated clinical trial. Specifically, develop a process for high frequency, bi-allelic CCR5 gene disruption with minimal manipulation and no negative impacts on cell engraftment or lineage potential, as well as assess the feasibility and reproducibility of the production process and in vivo safety of the final product.
CD34+ Cell Handling & Electroporation
Process Development
CD34+ cells were enriched from commercially-available, G-CSF mobilized hematopoietic progenitor cells collected by apheresis (HPC-A) followed by an overnight pre-stimulation regimen.5
- Stimulated CD34+ cells were resuspended in MaxCyte® Electroporation Buffer with either 0, 50, 75, or 150 mg/mL SB-728 mRNA.
- Cells were transferred to a CL1.1 processing assembly and electroporation performed using the recommended protocol on the MaxCyte GT.
- Electroporated cells were incubated for 20 minutes at 37°C, followed by overnight incubation at 30°C.11 Cells were then incubated an additional 24 hours at 37°C prior to analysis or cryopreservation.
Engineering Runs & In Vivo Turmogenicity Studies
- CD34+ HSPCs were isolated by apheresis from four G-CSF mobilized, healthy donors and used for GLP qualification runs.
- Stimulated CD34+ cells were electroporated as above using 150 mg/mL clinical-grade SB-728 mRNA.
- An aliquot of cells from each lot was cryopreserved and evaluated by MiSeq deep sequencing to determine the level of CCR5 disruption.
- Cryopreserved cells were thawed and cultured for 24 hours in media containing a cytokine cocktail prior to intravenous injection into NOD/SCID gamma (NSG) mice for a 20-week tumorigenicity/toxicity study.
Full methods for SB-728mR manufacturing, in vitro assays, sequencing, and in vivo safety studies are detailed in Mol. Ther. — Methods & Clinical Development, 3, 16067, 2016.
Results
Process Development Studies: 73% Bi-allelic CCR5 Disruption
By migrating from viral transduction to transient ZFN expression via mRNA electroporation, gene disruption rates could be maximized via mRNA concentration while minimizing off-target activity. Additionally, this eliminated the risk of adenoviral vector toxicity as noted in previous studies.5
Electroporated CD34+ enriched HSPCs showed viabilities of >90% with high frequency CCR5 gene modification during initial process development.12 Up to 73% bi-allelic gene disruption was seen, with disruption frequency correlating with the concentration of ZFN-encoding mRNA used (Table 1). The significance of such high-level bi-allelic disruption rates is highlighted by early studies that suggest biallelic CCR5 disruption is required for full R5-tropic HIV resistance and thus may be essential for therapeutic efficacy.2,3
In addition to high-frequency gene disruption, electroporation of HSPCs, using even the highest dose of ZFN-encoding mRNA, did not negatively impact the level of engraftment or alter the lineage potential of the cells observed during a 20-week in vivo NSG mouse study. This suggests that engineered HSPCs can act as a longterm source of multilineage, HIV-resistant immune cells.12
CCR5 Gene Disruption in CD34+ Cells Following mRNA Electroporation
| mRNA Concentration |
50 μg/mL SB-728mR |
150 μg/mL SB-728mR |
|---|---|---|
| Bulk Culture Disruption | 41 | 57 |
| Total CFU (#) | 192 | 198 |
| Modified CFU | 55% | 70% |
| Mono-allelic Disruption | 33% | 19% |
| Bi-allelic Disruption | 22% | 51% |
| Modified CFU with Bi-allelic Disruption | 40% | 73% |
Table 1. Electroporated HPSCs were evaluated via Cel-1% assay for overall CCR5 disruption in 2-week bulk cultures or individual colonies derived from 2-week CFU cultures via deep sequencing analysis. CFU = colony forming units.
Reproducible, Clinical-scale Manufacture of CCR5-disrupted HSPCs
Full qualification runs using the production process proposed for the anticipated clinical trial were performed to assess manufacturing feasibility, reproducibility, and donor-to-donor variability. Cells harvested from four healthy human donors were electroporated with 150 mg/mL of CCR5-targeted ZFN mRNA, twice the dose proposed for the clinical trial, thereby maximizing the ability to identify off-target ZFN activity and potential tumorigenic activity. Cell viability levels ranged from 82–92% with CCR5 disruption rates >50% as determined using deep sequencing for 3 of the 4 donors.12 Release testing for sterility, stability, and phenotypic analysis was conducted.
Engraftment Rates
| HSPC transferred | No EP Controls | 3 Qualification Runs |
|---|---|---|
| Mice with hCD45+ cells in blood | 78% | 88% |
| Mice with hCD45+ cells in bone marrow | 98% | 96% |
| CCR5 gene disruption in hCD45+ PBMC | 0% | 10-20% |
| CCR5 gene disruption in hCD45+ bone marrow | 0% | 18-31% |
Table 2. Mice received HSPC from three independent donors either following a qualification run or non-electroporatin (EP) control cells. The presence of CD45+ cells in peripheral blood mononuclear cells and bone marrow was assessed at 4-, 12- and 22-weeks.12 Results of 22-week analysis are reported here. CCR5 gene disruption in hCD45+ cells was determined using MiSeq analysis.
IND-Enabling Studies: Positive Safety Profile
In vivo adoptive transfer studies using immunodeficient, NSG mice were performed using cells from the clinical-scale qualification runs to determine if any undetected genomic modifications would lead to tumor formation or toxicity upon engraftment. In pre-IND discussions with the FDA, these studies were deemed sufficient for meeting preclinical safety expectations in regard to the anticipated clinical trial. NSG mice received 1x106 electroporated CD34+ cells (equivalent to a full human dose), or no electroporation cells from the same donors. At the time of engraftment, CCR5 disruption rates ranged from 54 - 67%. Engraftment rates of the engineered HSPCs in the bone marrow and blood were similar to those of no electroporated cells over the 22-week study (Table 2). No evidence of toxicity or tumorigenicity was noted demonstrating product safety.
Initiation of Human Clinical Trial
The bi-allelic disruption efficiency, manufacturability, reproducibility, and general safety demonstrated in these studies supported initiation of a clinical trial to assess the feasibility and safety in patients with HIV-1 (Clinical Trial #: NCT02500849).
Conclusion
These studies demonstrate how MaxCyte non-viral cell engineering technology provides the required efficiency, viability, and safety to attain the necessary therapeutic index, as well as the scalability and regulatory-compliance to rapidly navigate clinical development — all while simultaneously eliminating the cost, complexity, inconsistency, and potential toxicities of viral vectors. While this application note highlights the success of CCR5 disruption in HSPCs as a treatment for HIV, this non-viral gene editing approach has been employed for the treatment of a growing number of indications ranging from monogenic disorders such as sickle cell disease to cancer.
References
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- Tebas P, Stein DS, Tang WW, Frank I et al. Gene editing of CCR5 in autologous CD4 T cells of persons infected with HIV. (2014) N Engl J Med 370: 901–910.
- Liu R, Paxton WA, Choe S, Ceradini D, Martin SR, Horuk R, MacDonald ME, Stuhlmann H, Koup RA, Landau. Homozygous defect in HIV-1 coreceptor accounts for resistance of some multiply-exposed individuals to HIV-1 infection. (1996) Cell 86: 367–377.
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- Doyon Y, Choi VM, Xia DF, Vo TD, Gregory PD, Holmes MC. Transient cold shock enhances zinc-finger nuclease-mediated gene disruption. (2010) Nat Methods 7: 459–460.
- DiGiusto DL, Cannon PM, Holmes MC, Li L, Rao A, Wang J, Lee G, Gregory PD, Kim KA, Hayward SB, Meyer K, Exline C, Lopez E, Henley J, Gonzalez N, Bedell V, Stan R, Zaia JA. Preclinical development and qualification of ZFNmediated CCR5 disruption in human hematopoietic stem/progenitor cells. (2016) Mol. Ther. — Methods & Clinical Development, 3, 16067.
