Children with B-cell acute lymphoblastic leukemia (B-ALL) are increasingly treated with immunotherapeutic agents such as blinatumomab and inotuzumab. Thus, those who present in relapse for chimeric antigen receptor (CAR) T-cell therapy are likely to have prior exposure to these other immunotherapies. The effect of blinatumomab and inotuzumab exposure on cellular therapy, particularly, cell collection and the composition of apheresis products, is unknown. In this study, we analyzed the apheresis composition of 113 pediatric and young adult patients with relapsed or refractory B-cell malignancies across three CART-cell trials spanning from 2014 to 2024 and compared patients who did or did not have exposure to blinatumomab or inotuzumab prior to cell collection. We found no association between blinatumomab exposure and differences in apheresis composition, including total nucleated cell (TNC) yield, CD3 counts, CD56 counts, or CD4:CD8 ratio of CD45+ cells. The apheresis products from patients with inotuzumab exposure had lower CD4:CD8 ratios compared to those of patients without inotuzumab exposure. There were no other notable differences in T-cell phenotype or markers of exhaustion among the subset of samples with extended flow cytometry panels. With the exception of lower CD4:CD8 ratios in patients with prior inotuzumab, the comparable apheresis yield and composition observed after immunotherapy exposure is a reassuring finding, particularly in light increased use of upfront blinatumomab for B-ALL.

Abstract Background Chimeric antigen receptor (CAR) T-cell therapy has transformed the treatment landscape for many hematological malignancies. However, high relapse rates and limited accessibility remain significant challenges. We developed a 3-day streamlined process to address these limitations. Methods T-cells were isolated from whole blood collected from healthy donors via an automated density gradient separation that included a T-isopure™ antibody cocktail, which isolates T-cells through negative selection. T-cells were activated then transduced with a lentiviral vector encoding a trispecific CAR. T-cells were cultured in G-Rex vessels and harvested at day 3 or day 7 for analysis. CAR expression and T-cell phenotype were assessed by flow cytometry and gene expression analysis. Functional activity was evaluated by measuring cytotoxicity and cytokine secretion following co-culture with target cell lines. Results The T-isopure isolation enriched CD3+ T-cells in whole blood from 18.9% to 88.5% of CD45+ cells, with a mean recovery of 40.6%. RBCs were depleted with ≥ 99% efficiency, with monocytes and NK cells comprising the bulk of remaining CD45+ cells. The 3-day manufacturing process produced T-cells with > 95% viability, 53% transduction efficiency, and vector copy number < 3 copies/cell. Phenotypic analysis revealed a high proportion of stem/central memory T-cells at both timepoints, with no significant differences observed. Cytotoxicity assays demonstrated strong and sustained killing of NALM6 tumor cells, comparable between both products. Gene expression profiling indicated that day 3 products were less differentiated, exhibiting a memory-like phenotype and reduced inflammatory signaling, further supported by protein analysis of culture supernatants. Conclusion This study establishes a rapid, GMP-compliant method for manufacturing polyfunctional, CAR T-cells directly from whole blood. The workflow outlined here achieved a potent, phenotypically favorable CAR T-cell product without compromising viability or cytotoxic function. Compared to the standard 7-day method, the 3-day approach resulted in expression of genes associated with a more stem-like phenotype while reducing manufacturing time and cost. This method may provide a practical alternative for decentralized CAR T-cell manufacturing, particularly in resource-limited settings.

Chimeric antigen receptor (CAR) T-cells have become a standard therapy for many hematologic malignancies. While many CAR T-cells are being produced commercially and are in clinical trials, more CAR T-cells are being developed, particularly for the treatment of patients with solid tumors. Unlike most other therapies used to treat cancer and hematologic malignancies, CAR T-cells are biological agents which are most often produced from a patient’s own T-cells. A critical part of the development of new CAR T-cells is identifying the characteristics that are responsible for their potency, toxicity, and clinical effectiveness which often vary among patients. We describe factors responsible for CAR T-cell potency, ongoing work focused on understanding critical characteristics of CAR T-cells contributing to their potency and safety, and ways to improve their potency. Since the factors affecting CAR T-cell potency are dependent on their structure and manufacturing methods, the nature of CAR vectors and CAR T-cell manufacturing methods are also reviewed. When assessing CAR T-cell potency in vitro studies provide valuable information, but it is also important to use data obtained from early clinical trials of specific CAR T-cells to advance the understanding of factors and characteristics that contribute to CAR T-cell potency, safety and clinical effectiveness.

Fostamatinib had 46% durable response, with 73% steroid reduction, in this multicentre retrospective study of refractory wAIHA/ES. Hypertension, gastrointestinal (GI) distress and neutropenia occurred in 23%. Only one patient required drug discontinuation and one patient dose reduction. In an eight-patient subset, migration inhibitory factor (MIF) levels, naïve (CD62Lhi) T-cell and HLA-DRhi monocyte subsets each correlated with treatment response.

Introduction Despite success in relapsed/refractory (r/r) B-cell acute lymphoblastic leukemia (B-ALL), chimeric antigen receptor (CAR) T-cell immunotherapies have had limited efficacy in patients with r/r acute myeloid leukemia (AML). We observed only transient expansion of CAR T-cells in 9 of 19 children, adolescents, and young adults infused in CD33-targeted CAR T-cells (CD33CART) in our recently completed phase I clinical trial (NCT03971799), with complete response occurring in only 2 subjects. A potential etiology of lack of response to CD33CART is poor T-cell fitness or inhibitory immune cells within the collected leukapheresis products. Objective To compare leukapheresis products from patients with r/r AML or B-ALL to identify T-cell or myeloid cell features potentially associated with suboptimal CAR T-cell function. Methods Cryopreserved whole apheresis products from 14 patients enrolled (12 infused) on the CD33CART trial, 7 patients with B-ALL treated on Phase I CAR T-cell trials, and 4 healthy donors (HDs) were compared. Samples were analyzed via multiparameter flow cytometry for markers of T-cell differentiation, activation, and exhaustion and myeloid/monocytic cell subsets. Data were analyzed using FlowJo. Results Five of 12 infused apheresis products from patients with AML demonstrated in vivo CAR T-cell expansion, including one subject with CD33CART-induced complete response with incomplete count recovery (CRi). Among patients with B-ALL, all 7 proceeded to CAR T-cell infusion and 4 achieved CR.Overall, T-cell differentiation phenotypes and CD4:CD8 ratios did not differ amongst AML-, ALL-, or HD-origin apheresis products. (Fig 1) CD62L expression was notably higher in AML-origin T cell products, specifically in the T-effector population, in which CD62L loss is associated with enhanced lytic activity (45.5% of Teff in AML vs 27.6% in B-ALL; p=0.016). Expression of the CD39 exhaustion marker was higher in AML samples (11.8% AML vs 3.9% in ALL; p=0.015), while other exhaustion markers did not differ. CD39, CD69, LAG-3, and TIM3 expression was higher in AML-origin T-cell products compared with HD. CD4+ CD25+CD127dim T-regulatory cells did not vary between groups, although a higher proportions of Tregs was detected in a subset of patients with AML.(Fig 2)While AML-origin products had lower proportion of classical monocytes compared to those from HDs (72% vs 91%; p=0.046), no differences in monocyte subsets between B-ALL and AML groups were detected. (Fig 3) Conclusion Although leukapheresis products exhibited heterogeneity across patients, few differences in T-cell or monocyte phenotypes were seen in AML- versus B-ALL-origin samples. A trend towards increased T-cell exhaustion in AML patient-origin products was observed and merits evaluation in larger studies to understand potential contribution to poor CAR T-cell functionality in patients with r/r AML.

Transduction efficiency (TE) is a key measure of CAR T-cell manufacturing quality, however its variability and relationship to the final product phenotype remain incompletely understood. In this retrospective study, we evaluated TE across 204 GMP-manufactured CAR T-cell products, integrating manufacturing parameters, starting material composition, and immunophenotypic and transcriptomic profiles of final products. Multivariable linear regression and protocol-specific correlation analyses were performed to account for manufacturing-related confounding. TE variability was primarily associated with integrated manufacturing workflows and starting material composition. The CD4CD8-Prodigy-TransAct workflow was associated with higher TE compared with the CD4CD8-Bag-Dynabeads workflow, although contribution of individual manufacturing parameters could not be independently determined due to protocol-specific interdependencies. Within PBMC-based manufacturing, monocyte abundance was negatively associated with TE, suggesting protocol-dependent contributions of starting material composition. Higher TE was associated with a less differentiated T-cell phenotype, including lower effector memory T-cell (TEM) frequencies, particularly within CD4+CAR+ compartment. Transcriptomic analyses further identified associations between TE and proliferation- and oxidative phosphorylation-associated gene-expression signatures; however, these findings were exploratory and may reflect protocol-related differences. Collectively, TE variability was associated primarily with integrated manufacturing workflows, providing a real-world framework for understanding CAR T-cell manufacturing performance.

BACKGROUND:Accurate prediction of peripheral blood stem cell (PBSC) yield is essential for optimizing hematopoietic stem cell collections. Traditional CD34+ prediction formulas often overlook the ongoing recruitment of CD34+ cells during the collection procedure. This study evaluated whether a single intra-apheresis CD34+ measurement could improve yield prediction compared to existing prediction models. METHODS:We retrospectively analyzed 150 PBSC collections from 123 autologous and allogenic donors. A single intraprocedural CD34+ measurement and final collection yield were obtained, and univariable and multivariable linear regressions were developed to assess predictive accuracy. Model stability and generalizability were tested via bootstrap resampling and cross-validation. Subgroup analyses by age, sex, weight, diagnosis, and sampling time were performed to confirm robustness. We also compared our results against published pre-apheresis formulas. RESULTS:Intra-apheresis CD34+ counts showed a strong correlation with the final yield (r = 0.97; R2 = 0.95). Subgroup analyses across donor demographics and clinical factors yielded similarly high R2 values (0.91-0.99). Multivariable analysis identified intra-apheresis CD34+ measurement as the primary contributor (β = 0.67, p <.0001). Later sampling (≥1.20 of total blood volumes processed) reduced error metrics relative to earlier draws. Compared to pre-apheresis models, the intra-apheresis approach achieved higher R2 and lower error metrics. DISCUSSION:A single intra-apheresis CD34+ measurement, particularly at or beyond 1.20 processed blood volumes, offers a reliable, real-time predictor of final PBSC yield. This intraprocedural approach outperforms published pre-apheresis formulas, enabling more efficient collection and reduced donor burden across diverse clinical scenarios.

Multiantigen targeting chimeric antigen receptor (CAR) T cells have emerged as a strategy to mitigate antigen escape observed after single antigen targeting therapy. Our initial experience with a bivalent CD19.22.BBζ CAR T-cell construct in children, adolescents and young adults (CAYA) with B-cell acute lymphoblastic leukemia (B-ALL) demonstrated limitations in CD22 recognition, but a tolerable safety profile and efficacy prompted further evaluation. This trial enrolled patients between the ages of 3-39 with relapsed/refractory B-ALL. Following dose-escalation, patients who enrolled at the recommended phase 2 dose (RP2D) of 3×106 transduced CAR T cells/kg constitute this report. 30 CAYA were treated at the RP2D; 28 with B-ALL and 2 with Burkitt lymphoma. Across patients with B-ALL, 20 (71.4%) patients developed cytokine release syndrome (CRS); only 2 (10%) were grade >3. Grade 3 immune effector cell-associated neurotoxicity syndrome (ICANS) occurred in 3 (10.7%) patients; there were no cases of immune effector cell-associated hemophagocytic lymphohistiocytosis-like syndrome. Following a protocol amendment to evaluate the efficacy of siltuximab as a first-line treatment of CRS, one patient received siltuximab with full resolution of CRS after two doses without needing additional anti-cytokine-directed therapies. A measurable residual disease-negative complete remission (CR) was attained in 25 (89.3%) patients, including 6 who had neither CRS nor ICANS. 23 patients (82.1%) proceeded directly to hematopoietic stem cell transplant (HSCT) following CAR T-cell infusion within a median of 51 days (range, 45-68 days), supporting the utility of this construct as a bridge to HSCT. All three non-responders had persistent non-central nervous system (CNS) extramedullary disease (EMD), although four of seven patients with non-CNS EMD achieved a CR. Median relapse-free survival among the 25 patients achieving CR was not reached, and the median overall survival for all 28 patients was 34 months (95% CI 17 to not estimable) from infusion. This extended experience demonstrates that CD19.22.BBζ CAR T-cell therapy is safe and clinically active, particularly as a bridge to HSCT. Non-response was confined to patients with non-CNS EMD, highlighting the persistent challenge of effectively targeting EMD and informing the design of future CAR constructs.Trial registration numberNCT03448393.

BACKGROUND:Hematopoietic progenitor cells (HPCs) and mononuclear cells (MNCs) are critical components of cell-based therapies, including bone marrow transplantation and regenerative treatments. Evaluation of the characteristics of these products during collection, storage, and transport is essential for maintaining cell viability and functionality. In this study, we evaluated the functional and molecular stability of samples collected for the evaluation of fresh HPC and MNC products. The samples stored at 4 °C for up to 4 days and were evaluated using white blood cell (WBC) counts, flow cytometry, and bulk RNA sequencing (RNA-seq) across five time points. METHODS:HPC samples from seven products (June-December 2022) and MNC samples from six products (October 2022-August 2023) were analyzed on days 0 through 4 after collection. WBC counts were measured, and viability was assessed using 7-AAD staining and flow cytometry. HPC samples were stained with antibodies against CD34, CD3, CD19, CD56, CD14, CD16, CD15, and CD45, while MNC samples were stained with antibodies directed to CD3, CD4, CD8, CD19, CD56, CD14, CD16, CD15, and CD45. Total RNA was isolated from each sample and subjected to bulk RNA-seq to assess transcriptomic changes during storage. RESULTS:While WBC counts varied between products, no significant differences were observed across time points within individual products. Flow cytometry markers remained relatively stable over time in both HPC and MNC samples, although greater variability was observed in HPCs. A modest decrease in lymphocyte percentages was noted at later time points, primarily driven by a reduction in CD3+ cells; however, these changes were not statistically significant. Cell viability declined significantly over time within individual products and showed inter-product variability. RNA-seq analysis revealed stable gene expression profiles in MNC samples across all time points. In contrast, HPC samples exhibited notable transcriptomic changes as early as day 1 of storage at 4 °C, indicating greater molecular instability. CONCLUSION:WBC counts and flow cytometry markers remain stable for up to 3 days in samples collected from fresh HPC and MNC products when stored at 4 °C, although cell viability progressively declines. However, RNA-seq data reveal early transcriptomic changes in HPC samples, suggesting that immediate evaluation of these samples is critical to preserve their molecular integrity and functionality. These findings support the feasibility of delayed phenotypic analysis but emphasize the need for prompt molecular assays in HPC-based applications.

Although CAR T cell therapy is increasingly used to treat relapsed B cell acute lymphoblastic leukemia (ALL), 20%- 30% of patients do not respond, and few clinical predictors of response have been established, especially in the pediatric population. A deeper analysis of CAR T cell infusion products, along with the apheresis product used as the starting material for CAR T cell manufacturing, provides valuable insights for predicting clinical outcomes. We analyzed infusion products and CD4/8-selected T cell starting materials from pediatric and young adult patients on a single-center study with relapsed/refractory B cell ALL who were undergoing treatment with CD22 CAR T cells and evaluated differences between T cells from responders and non-responders (NCT023215612). We found that CAR T cells from non-responders had a more differentiated T cell phenotype and over-expressed genes associated with cytotoxicity and exhaustion compared with those of responders. Furthermore, we found that these differences could be tracked back to the apheresis materials prior to CAR T cell manufacturing. Using flow cytometry-based immunophenotypic markers, we developed a scoring system that distinguished non-responders based on T cell phenotype at the time of apheresis. These findings can help inform outcomes for patients and providers as well as provide insights into targeted manufacturing changes to optimize CAR T cell efficacy.

Background: Chimeric antigen receptor T (CAR-T) cells have significantly advanced the treatment of cancers such as leukemia and lymphoma. Traditionally, T cells are collected from patients through leukapheresis, an expensive and potentially invasive process that requires specialized equipment and trained personnel. Although whole blood collections are much more technically straightforward, whole blood starting material has not been widely utilized for clinical CAR-T cell manufacturing, in part due to lack of manufacturing processes designed for use in a good manufacturing practice (GMP) environment. Collecting cellular starting material from whole blood without leukapheresis could reduce manufacturing complexity and cost, thereby improving accessibility to CAR-T cell therapy. Methods: Whole blood samples were collected from eight healthy donors and one pediatric B-cell acute lymphoblastic leukemia (B-ALL) patient. These samples were processed using the Sepax C-Pro (Cytiva) instrument to isolate mononuclear cells (MNCs) via density gradient separation. CAR-T cells were then manufactured from the isolated MNCs using a GMP-compliant 7-day protocol, whereby T cells were activated with anti-CD3 and IL-2, transduced with GMP lentiviral vector encoding a CD19/CD22 bispecific CAR, and expanded in gas permeable cell culture bags. The resulting CAR-T cells were then evaluated for their phenotypic and functional properties using flow cytometry, cytokine release and cytotoxicity assays. Results: From an average 77.7 mL of whole blood from healthy donors (range = 29-96 mL), we isolated an average of 42.2 x 106 CD3+T cells (range 7.3-63.0) postprocessing. CAR-T cell cultures were initiated from thaw using 1-10 x 106 starting CD3+ T cells, yielding a median T cell number of 105 x 106 cells on day 7 (range 61-188 x 106). We observed 66 f 11% mean transduction efficiency and produced a mean of 77.4 x 106 transduced CAR-T cells (range 30.8-143.5 x 106). Similar results were obtained when using a blood sample (28mL) obtained from a patient with relapsed B-ALL who had received recent chemotherapy. Conclusions: Therapeutically relevant doses of CD19/CD22 CAR-T cells can be successfully manufactured from whole blood. On average, 80 mL of whole blood yields enough CAR-T cells to create a single dose for a pediatric patient (50 kg) at a dosage of 1 x 106 CAR-T cells/kg. For larger patients, scaling up is straightforward by collecting a larger blood volume. This method also demonstrates a cost-effective approach to T cell activation and expansion which, alongside a more straightforward collection of whole blood, makes it more widely accessible especially for middle- and low-income countries. By reducing costs and labor, this strategy has the potential to significantly expand global access to CAR-T cell therapy. Published by Elsevier Inc. on behalf of International Society for Cell & Gene Therapy. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/)

Background Toll-like receptor (TLR) agonists and radiation therapy hold promise for cancer immunotherapy. We conducted a phase I/II trial combining topical imiquimod (IMQ, a TLR-7 agonist) and local radiotherapy (RT) in patients with metastatic breast cancer accompanied by longitudinal transcriptional analysis of tumor biopsies.Methods The primary objective of the trial (NCT01421017) was to assess systemic responses by immune-related response criteria (irRC) after an 8-week cycle of topical IMQ and concurrent local RT (cohort 1). An amendment to the trial added two cohorts, both received one dose of cyclophosphamide (CTX) administered 1 week before study treatment initiation, IMQ/RT/CTX (cohort 2) and RT/CTX control (cohort 3). Cutaneous metastases were prospectively assigned to treatment with IMQ and RT (area A) or IMQ alone (area B). Secondary objectives were safety (Common Terminology Criteria for Adverse Events criteria) and local response in skin metastases. In all IMQ cohorts, tumors were biopsied before treatment and at 2 and 3 weeks.Results 31 patients were enrolled (n=12, n=12, and n=7, in cohort 1, 2, and 3, respectively), with 4 out of 24 patients in the IMQ cohorts showing systemic tumor responses (two complete responses (CR) and two partial responses (PR)). No objective responses were observed in the seven patients enrolled in the control arm (RT alone). The treatment was well-tolerated, no grade 4–5 treatment-related adverse events occurred and grade 3 AEs were manageable (anemia, local pain, and local ulceration, n=1 each). Local objective responses were observed in 19/24 (9 CR and 10 PR) and 5/24 (5 PR) in areas treated with combined IMQ-RT and IMQ alone, respectively (p<0.001). All 24 patients treated with IMQ underwent serial biopsies, and 84 samples yielded sufficient material for transcriptional analyses. These revealed that the presence of a T-helper 1 functional orientation of the tumor microenvironment paralleled by the downregulation of DNA-repair genes was associated with CR after IMQ+RT, but not after IMQ alone. No post-treatment activation of immune-effector functions was observed in stable and progressing lesions.Conclusions Our findings support the safety and clinical efficacy of combining topical IMQ with local RT for recurrent breast cancer, with evidence of local and occasional systemic antitumor activity.Trial registration number NCT01421017.

This review explores recent advances in the characteristics and manufacturing of CAR T-cell products. Traditional potency assays have been designed based on well-established CAR T-cell functionalities. However, the advent of innovative tools and methodologies has revealed a broader spectrum of important CAR T-cell characteristics that correlate with function. Furthermore, as manufacturing strategies continue to evolve, conventional potency assays may no longer fully capture the complexity of these products. Therefore, it is essential to examine these emerging characteristics and manufacturing approaches and consider the development of tailored potency assays to ensure products are fully characterized.

IntroductionEmerging data links characteristics of the apheresis product with outcomes following chimeric antigen receptor (CAR) T-cell therapy, but data on predictive markers in the pediatric B-cell acute lymphoblastic leukemia (B-ALL) population are limited. Given the ability to modify manufacturing and pre-collection therapies to optimize CAR T-cell functionality, identifying key biomarkers at the time of apheresis is imperative to guide decision making and improve outcomes.MethodsWe retrospectively analyzed CD4/CD8-selected apheresis (T-cells) from children and young adults who were treated with CD22 CAR T-cells for B-ALL (NCT02315612). Analysis included flow cytometry for T-cell immunophenotype, activation, and exhaustion markers, Seahorse metabolic profiling, and RNA sequencing. We compared patients who achieved a complete response (CR) with those who had partial response, progressive, or stable disease (NR). Additional analysis compared patient T-cells to those of healthy donors. Results were analyzed using Mann-Whitney tests. Differential gene expression analysis identified genes with >2x fold change.ResultsAcross 24 patient apheresis samples, 16 (67%) achieved CR and 8 (33%) did not (NR). Demographics, with exception of disease burden, were comparable. As expected, median pre-infusion disease burden was lower in patients with CR: 28% (r: 0.04-98.2%) vs 59% (r: 0.33-97.9) in NR (p=0.18).Analysis by UMAP and FlowSOM distinguished NR from CR by absence of activated, naive CD4 cells, and presence of CD4 effector cells. (Fig 1) Manual gating confirmed that patients with NR had lower CD4/CD8 ratios, fewer naïve (CD45RA+) CD4 T-cells, lower expression of CD28 and CD127 (IL-7R) (activation markers), and higher CD69. (Fig 2) RNA sequencing similarly showed lower IL-7R expression.Among patients with M3 marrow (>25% blasts) (n=15), those with NR (n=6) still had significantly fewer CD127+, CD8+CD28+, and CD4+CD45RA+ cells and more CD69+ cells compared to those with CR. Specifically, having <50% of CD4 cells expressing CD127 fully separated NR from CR in this M3 cohort.Compared to healthy donors, patients had more TIM3, LAG3, and CD39. However, those with CR resembled healthy donors in all markers that distinguished them from NR (Fig 3). Additionally, while healthy donors had higher ATP production and respiratory capacity and less glycolysis compared with B-ALL patients, metabolic characteristics did not distinguish CR from NR.ConclusionBased on our analysis, starting material characteristics in B-ALL can predict NR, using a flow cytometric assay alone. This information can provide real-time guidance to patients, families, and providers on expectations. Additionally, CD4+CD127+ cells in the starting material appear to be necessary to overcome high disease burden, suggesting the importance of optimized manufacturing to expand this favorable cell subset.

Background & AimMutations in CYBB gene disrupt gp91phox expression causing X-CGD, an inherited immune deficiency disorder of granulocytes with impaired NADPH oxidase activity for pathogen killing. We address a common missense mutation that accounts for ∼6% of our clinical X-CGD patient cohort. We developed a process to precisely correct the mutation at genome level in autologous HSPCs using pioneering CRISPR base-editing technology for ex vivo gene therapy.Methods, Results & ConclusionA total of 3 performance qualifications were completed using the post-thawed CD34+ cells enriched from mobilized apheresis products from 2 healthy donors (to knock down gp91phox expression) and 1 patient (to restore gp91phox expression). Thawed CD34+ cells were pre-stimulated for up to 48 hours before electroporation with Adenosine base editor (ABE) and single guide RNA using MaxCyte GT device. Final products (∼48hr post-EP) were cryopreserved after submitting FN assays (cell count, viability, purity, sterility, mycoplasma, endotoxin, base editing rate by deep sequencing, gp91 expression and DHR function in granulocytes differentiated in vitro from final product by flow cytometry). Cells electroporated at varying concentrations, 50-200x106/mL exhibited similar viability, purity and fold expansion in healthy donor final products. We observed slightly lower viability and purity in the patient sample which remained well within the range of acceptability (≥70%). Base editing data from HD cells showed high performance consistency when using 50-200x106cells/mL for electroporation (31.3-34.49%), and robust base editing rate with the larger CL1.1 processing assembly (52.42%). Base editing rate using the mutation-specific base editor/sgRNA was 64.97% correction, which demonstrated the process feasibility. Functionally, we observed in myeloid-differentiated CD34+ successful disruption of gp91 expression (74.8% to 32.7%) and DHR function (74.6% to 28.1%) in HD naïve compared to BE-products. In patient cells, restoration of gp91 expression (0.47% to 61.9%) and DHR function (from 1.6% to 49.3%) was observed in naïve compared with BE-CGD products. Furthermore, cryopreservation did not demonstrate any detrimental effects on final product viability, purity and potency. In summary, the process for GMP manufacturing of base-editing autologous HSPC was successfully validated and generated products that passed all release criteria. We anticipate the start of 1st approved base editing clinical trial (IND 28925) for XCGD by Mid-2024.

Background aims: Accurate assessment of cell viability is crucial in cellular product manufacturing, yet selecting the appropriate viability assay presents challenges due to various factors. This study compares and evaluates different viability assays on fresh and cryopreserved cellular products, including peripheral blood stem cell (PBSC) and peripheral blood mononuclear cell (PBMC) apheresis products, purified PBMCs and cultured chimeric antigen receptor and T-cell receptor-engineered T-cell products. Methods: Viability assays, including manual Trypan Blue exclusion, flow cytometry-based assays using 7-aminoactinomycin D (7-AAD) or propidium iodide (PI) direct staining or cell surface marker staining in conjunction with 7-AAD, Cellometer (Nexcelom Bioscience LLC, Lawrence, MA, USA) Acridine Orange/PI staining and Vi-CELL BLU Cell Viability Analyzer (Beckman Coulter, Inc, Brea, CA, USA), were evaluated. A viability standard was established using live and dead cell mixtures to assess the accuracy of these assays. Furthermore, precision assessment was conducted to determine the reproducibility of the viability assays. Additionally, the viability of individual cell populations from cryopreserved PBSC and PBMC apheresis products was examined. Results: All methods provided accurate viability measurements and generated consistent and reproducible viability data. The assessed viability assays were demonstrated to be reliable alternatives when evaluating the viability of fresh cellular products. However, cryopreserved products exhibited variability among the tested assays. Additionally, analyzing the viability of each subset of the cryopreserved PBSC and PBMC apheresis products revealed that T cells and granulocytes were more susceptible to the freeze-thaw process, showing decreased viability. Conclusions: The study demonstrates the importance of careful assay selection, validation and standardization, particularly for assessing the viability of cryopreserved products. Given the complexity of cellular products, choosing a fit-for-purpose viability assay is essential. (c) 2023 Published by Elsevier Inc. on behalf of International Society for Cell & Gene Therapy. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/)

Background & AimMononuclear cell (MNC) apheresis products serve as the starting material of many cell therapies, and they can be further processed to yield purified peripheral blood mononuclear cells (PBMCs). This PBMC purification step eliminates excess red blood cells (RBCs) and platelets, since these can inhibit the culture and/or expansion of specific cell types of interest such as T cells, NK cells, or monocytes. The use of Ficoll density gradient separation efficiently separates PBMCs from RBCs, and this process has been adapted to the automated Sepax C-Pro device (Cytiva) for clinical manufacturing. In contrast to the Sepax, the Curate Cell Processing System uses microfluidic technology to separate PBMCs without the use of Ficoll.Methods, Results & ConclusionWe performed a series of side-by-side PBMC purifications using the Sepax and Curate systems with 6 healthy donor MNC apheresis products. The Curate microfluidic system yielded ≥75% recovery of total nucleated cells (average = 78.7 ± 2.7%), which was a 48% increase compared to Sepax, with a 77% increase in recovery of CD3+ T cells in particular. This improvement in recovery was accompanied by a significant decrease in both RBC and platelet contamination, with a 99.7 ± 0.1% reduction of platelets in the Curate compared to 90.9 ± 0.6% in the Sepax. Characterization of T cells following isolation showed an equivalent T cell differentiation state of naïve vs. effector cells between the two platforms, and current experiments are underway to evaluate performance of isolated PBMCs in a CAR-T manufacturing process. Given the improved recovery and purity of PBMCs isolated from the Curate microfluidic system, this technology demonstrates significant utility particularly for applications where large cell numbers and efficient cell recovery are desired.