Paper Details

ABSTRACT

 Chimeric antigen receptor (CAR) T-cell therapies have transformed the management of several hematologic malignancies and are increasingly being evaluated in additional immune-mediated indications. However, broader clinical implementation remains constrained by complex manufacturing workflows, reliance on leukapheresis starting material, end product variability, and cost. In parallel, emerging clinical experience highlights the importance of more precisely defining product attributes, including T-cell subset composition, that may influence persistence, potency, and clinical outcomes, with some subsets associated with clinical benefit (e.g., naïve and memory subsets) and others associated with less favorable outcomes (e.g., regulatory T cells). To address these challenges, we evaluated a semi-automated, GMP-compatible workflow designed to generate CAR-T cells from whole blood while reducing manual handling and supporting control of the final cellular composition.

 Whole blood was processed using the CTS™ Rotea™ system for PBMC isolation and staining with a GMP-compatible flow cytometry panel, followed by sorting on the Sony CGX10 Cell Isolation System. The panel was designed to purify naïve CD4 T cells and total CD8 T cells at a predefined CD4/CD8 ratio while depleting monocytes and regulatory T cells. Sorted cells were collected directly into TexMACS medium and seeded into G-Rex bioreactors for activation, lentiviral transduction on day 2, and expansion until day 7. At the end of culture, viability, CAR expression, CD4/CD8 ratio, and T-cell phenotype were assessed by flow cytometry. Functional activity was assessed using cytotoxicity assays after co-culture with target tumor cell lines.

 Compared with a manual Ficoll-based workflow, use of the CTS™ Rotea™ system for PBMC isolation and staining reduced hands-on processing time, and optimization of the process on the CTS™ Rotea™ allowed a reduction of approximately 85% of the sorting time, while maintaining post-processing purity and viability above 95%. The CGX10 sorting strategy enabled efficient removal of CD4^lo monocytes and CD25^hi regulatory T cells, enrichment of CD4^hiCD45RA^hi naïve CD4 T cells, and collection of total CD8 T cells at the targeted ratio, while minimizing reagent complexity. The CGX10 mixed-ratio sorting and buffer-exchange capabilities supported direct collection into culture medium (TexMACS Medium) for immediate transfer to G-Rex bioreactors.

By day 7, the process generated viable T-cell products with 91.3% ± 3.0 viability (n=7), a fold change of 28,5 ±5,7 (n=7), CAR expression of 36.9% ± 11.7 (n=7), and a CD4:CD8 ratio normalized to an average 1.4:1, with variation depending on the donors and activation technology used. Post expansion phenotypic analysis confirmed efficient removal of regulatory T cells and enrichment of less differentiated T cell populations. Cytotoxicity assays confirmed the effector function of the generated CAR T cells.

A preliminary cost analysis indicated that operating expenses (OPEX) for this workflow may be reduced compared with mainstream automated manufacturing platforms, with additional potential for capital-expenditure (CAPEX) optimization through scale-up efficiencies, increased process integration, and automation.

Together, these initial bench-scale data support the feasibility of combining automated cell preparation using instruments such as the CTS^TM Rotea^TM system with CGX10-based cell sorting to manufacture CAR-T cells in Grex device from whole blood as starting material using a rapid, scalable, and GMP-compatible process. This approach may help reduce the burden on patients and clinical teams at the time of starting material collection, improve control of the target product composition, and lower overall manufacturing cost. Additional phenotypic characterization, control of the CD4:CD8 ratio during expansion, potency testing, and in vivo evaluation will be important to refine clinically relevant product attributes and guide further process optimization toward translational and clinical development. At scale, integrated instrument communication and automation may further support manufacturing efficiency and cost reduction.

The authors would like to thank Mohamad Kassem and Jerome Cabeau from Thermo Fisher Scientific for the support provided on this work.