Cryopreserved leukapheresis enables scalable and distributed CAR-T manufacturing: a multi-platform comparative study

Fresh leukapheresis exhibits time-sensitive viability decay (24-72-hour transport window)^1,2,3. Current CAR-T production faces a paradoxical dilemma: While up to 33% of lymphoma patients are unable to receive treatment due to leukapheresis failure or rapid disease progression⁴, the field still critically depends on cells sourced from these same immunocompromised patients for most cryopreservation protocols. These failure rates directly correlate with therapeutic outcomes: B-cell acute lymphoblastic leukemia (B-ALL) sees 15–40% treatment failures, rising to over 50% in B-cell lymphoma⁵. Intensive preconditioning regimens exacerbate T-cell deterioration. Furthermore, both advancing age and altered physiological conditions contribute to a progressive decline in immune cell quantity and function⁶. These studies indicate that cell quality impacts cell therapy outcomes. PBMCs cryopreservation demonstrates 20-30-year immune cell preservation⁷. Thus early storage is critical—driving our cryopreservation optimization.

Frozen PBMCs have a longer useful life and a wider application paradigm in cell therapy. However, with the continuous development of sorting technology, the selection of raw materials for cell therapy is more and more inclined to be treated directly with leukapheresis collection^8,9, rather than using PBMCs. Traditional PBMC isolation via density gradient centrifugation (e.g., Ficoll) is susceptible to temperature sensitivity and dilution ratio during processing¹⁰, leading to significant cell loss, particularly of monocytes and stromal cells, which are critical for T-cell activation and immune function^11,12. Direct cryopreservation of leukapheresis collections can significantly alleviate these bottlenecks, preserving cellular diversity and cell yields. Nevertheless, critical knowledge gaps persist in the systemic evaluation of cryopreserved leukapheresis for industrial applications. Most studies focus solely on T-cell function¹³, neglecting non-T cell components and their cryotolerance¹⁴.

This study demonstrates that the integration of standardized and automated closed production technologies for cryopreserved leukapheresis, when applied as starting materials for cutting-edge CAR-T platforms, constitutes a pivotal advancement toward distributed manufacturing. Implementing these innovations into future CAR-T therapeutic frameworks will not only provide novel raw materials for personalized treatments but also drive the transition of industrialized production toward distributed models, thereby enhancing therapeutic efficacy and safety.

Systematic bibliometric analysis: research trends and key breakthroughs in cryopreserved leukocyte technology

A systematic search was conducted across PubMed using keywords: ” (“PBMC” OR “peripheral blood mononuclear cells”) AND (“cryopreservation” OR “freezing” OR “long-term storage”), (“leukapheresis” OR “leukopak”) AND (“cryopreservation” OR “freezing”) Inclusion criteria encompassed peer-reviewed studies (2010–2024) focusing on cryopreservation protocols, clinical outcomes, or process standardization. The search retrieved 285 studies on cryopreserved PBMCs and 64 studies on cryopreserved leukapheresis products, accounting for only 18.3% of total studies (Fig. 1A). From 2010 to 2024, annual publications on leukapheresis cryopreservation (2–8 papers) remained lower than those on PBMC cryopreservation (16–29 papers) (Fig. 1B). Notably, the surge in PBMC-related publications in 2016 coincided with the rising demand for CAR-T therapies. CiteSpace analysis of 349 publications revealed key research clusters, including “cellbanker”, “flow cytometry”, “biomonitoring”, “T cells”, and “CAR-T cells” (Fig. 1C-1D), highlighting major themes: cell banking, quality control, immune monitoring, T/B cell biology, cell expansion, and cancer therapeutics. Burst keyword analysis (Fig. 1E) demonstrated early focus on myeloid-derived suppressor cells and T cells, with later emphasis on PBMCs and whole blood, reflecting shifting research priorities. As a starting material for CAR-T products, leukapheresis offers advantages by reducing donor phlebotomy frequency and serving as a standardized, traceable raw material for off-the-shelf therapies. However, research on cryopreserved leukapheresis remains inadequate, lacking standardized protocols and post-thaw functional validation. Systematic evaluation of leukapheresis cryopreservation should address subpopulation stability, functionality, and integration with automated platforms to ensure reproducibility.

Breakthrough in closed automated process: standardization of CAR-T manufacturing using cryopreserved leukapheresis

The figure (Fig. 2) outlines the comprehensive process of developing a standardized cryopreservation protocol for leukapheresis products, followed by a comparability study, and finally, the different manufacturing processes for CAR-T cells. Initially, we embarked on developing the leukapheresis cryopreservation process to establish a standardized protocol for cryopreserved leukapheresis products. The primary objective of cryopreserved leukapheresis is to preserve leukocyte components, with current research focusing on optimizing T-cell quality post-thaw. While leukocyte cryopreservation technology is relatively mature, the critical challenge lies in mitigating the impact of non-target cellular impurities (e.g., residual red blood cells, platelets) in leukapheresis products on post-thaw T-cell viability, recovery, and subsequent CAR-T product quality. To address this, a centrifugation-based strategy was systematically implemented to remove non-cellular impurities. Furthermore, clinical-grade cryoprotectant CS10 (10% DMSO) was selected to minimize erythrocyte volume interference with DMSO concentration, ensuring consistent cryoprotection efficacy.

Key parameters for leukocyte processing and cryopreservation are summarized in Table 1. The median cell concentration decreased progressively from 5.09–9.71 × 10⁷ cells/ml at initial leukapheresis to 4.06–5.12 × 10⁷ cells/ml pre-cryopreservation and further to 3.49–4.67 × 10⁷ cells/ml post-cryopreservation. Viability remained high throughout, with initial values of 99.2–99.5%, declining slightly to 94.0–96.15% pre-cryopreservation and 90.9–97.0% post-thaw. The CD3 + T lymphocyte proportion showed minimal variation, ranging from 43.82 to 56.31% initially to 41.19–56.45% pre-cryopreservation and 42.01–51.21% post-thaw, indicating no significant loss of T cells during processing. Formulation efficiency was optimized using a closed-system automated platform, reducing processing times to 43–108 min. Cryopreservation parameters included a target cell concentration of ~ 5 × 10⁷ cells/ml to accommodate high-density requirements. Post-thaw viability (90.9–97.0%) and CD3 + purity (42.01–51.21%) confirmed effective cryoprotection and minimal cellular damage. Initial leukapheresis specifications included a leukocyte volume of ~ 1 ml per 1 × 10⁹ cells, hematocrit levels of 5–10%, and residual volume retention (3 ml per 1 × 10⁹ cells) to ensure ≥ 7.5% DMSO concentration in the cryomedium. These refinements underscored robust process reliability and consistency.

Optimized cryopreservation concentration: A target concentration range of 5 × 10⁷–8 × 10⁷ cells/ml was established, with a formulation volume of 20 ml/bag, ensuring ≥ 1 × 10⁹ cells per bag as a critical quality attribute (CQA). Time-sensitive freezing protocol: The interval from cryoprotectant addition to controlled-rate freezing initiation was strictly limited to ≤ 120 min (Table 2), validated using the Thermo Profile 4 system to prevent ice crystal formation and ensure post-thaw viability ≥ 90%.

Comprehensive functional profiling and comparative analysis of cryopreserved leukapheresis products versus PBMCs

To investigate the impact of the freezing process on the performance of cryopreserved leukapheresis products versus PBMCs, we conducted a comprehensive functional profiling and comparative analysis. Starting with an equivalent amount of leukapheresis material (Fig. 3A), we examined the phenotypic differences between fresh and cryopreserved samples.

Our analysis revealed a statistically significant difference in the proportion of major immune cells, specifically lymphocytes, between fresh and cryopreserved leukapheresis products compared to cryopreserved PBMCs. Fresh and cryopreserved leukapheresis products exhibited a higher percentage of lymphocytes (Fresh-LEUK: 68.68 ± 1.78%, Cryo-LEUK: 66.59 ± 2.64%) compared to cryopreserved PBMCs (Cryo-PBMC: 52.20 ± 9.29%, p < 0.05) (Fig. 3B). In contrast, the proportions of neutrophils and monocytes were similar across all groups, with no statistically significant differences observed.

We further compared three key subsets within the lymphocyte population, including T, B, and NK cells. Our results indicated that there were no significant differences in the proportions of these subsets between the groups (Fig. 3C). The higher lymphocyte count in cryopreserved leukapheresis products was primarily attributed to an increased proportion of T cells, which may be advantageous for T cell-based therapies such as CAR-T. As a critical raw material for CAR-T manufacturing, we investigated the phenotypic and functional characteristics of T cells. The ratios of CD4 + and CD8 + T cells, as well as their functional subsets (Tn, TCM, TEM, and Teff), were comparable between thawed leukapheresis products and PBMCs (Fig. 3D and 3E). This suggests that the freezing process did not significantly alter the T cell composition or functional characteristics.

We compared the viability of cells at various stages of CAR-T manufacturing. Post-thaw recovery and T cell sorting resulted in lower viability for both cryopreserved leukapheresis products and PBMCs compared to fresh samples. Specifically, the viability of cryopreserved samples was significantly lower than fresh samples at the pre-separation stage (Fresh-LEUK: 99.04 ± 0.30% vs. Cryo-LEUK: 90.99 ± 0.86%, p < 0.0001; Fresh-LEUK: 99.04 ± 0.30% vs. Cryo-PBMC: 93.47 ± 1.95%, p < 0.01) and post-separation stage (Fresh-LEUK: 98.78 ± 0.17% vs. Cryo-LEUK: 94.06 ± 0.87%, p < 0.001; Fresh-LEUK: 98.78 ± 0.17% vs. Cryo-PBMC: 93.97 ± 1.42%, p < 0.01). However, before transduction, the viability of cryopreserved leukapheresis products was restored to levels comparable to fresh samples (Fresh-LEUK: 93.34 ± 0.78% vs. Cryo-LEUK: 93.88 ± 0.56%, p > 0.05; Fresh-LEUK: 93.34 ± 0.78% vs. Cryo-PBMC: 90.70 ± 1.01%, p > 0.05) (Fig. 3F).

We then focused on the recovery of cryopreserved leukapheresis products versus cryopreserved PBMCs after thawing. No statistically significant difference was observed in the recovery rates between the two groups (Fig. 3G). However, starting with an equivalent amount of leukapheresis material, cryopreserved leukapheresis products significantly preserved a greater number of leukocytes (97.5% ± 4.4%) compared to cryopreserved PBMCs (70.0% ± 6.2%, p < 0.01) (Fig. 3H). Consequently, this led to a higher yield of T cells in cryopreserved leukapheresis products (34.2% ± 8.6%) compared to cryopreserved PBMCs (21.7% ± 8.6%, p < 0.01) (Fig. 3I).

Cross-platform adaptability verification: cryopreserved leukapheresis as enablers of novel CAR-T production frameworks

To investigate the feasibility of using cryopreserved leukapheresis as the initial raw material for conventional CAR-T manufacturing with the non-viral vector Piggy Bac transposon system, we compared the cell viability during the culture process (Fig. 4A) and continuous growth (Fig. 4B), finding no statistical difference between cryopreserved and fresh leukapheresis. Simultaneously, we explored the effects of cryopreserved and fresh leukapheresis on cell phenotype, cytotoxicity, and exhausted phenotype during CAR-T manufacturing. The study found no statistical difference in the proportions of Tn, TCM, TEM, and Teff between cryopreserved and fresh raw materials (Fig. 4C). For CAR-T, cryopreserved leukapheresis had no statistical difference from fresh one (Fig. 4C). Through the analysis of exhaustion functional markers (PD1+, LAG3+, and TIM3+) of CD3 + T cells on Day 9, we found no statistical difference between cryopreserved and fresh leukapheresis either (Fig. 4D). Finally, we conducted an assay of cellular cytotoxicity with effector: target (E: T) ratios of 2:1, 4:1, and found no statistical difference in the cytotoxicity effect between the two materials (Fig. 4E). Overall, cryopreserved leukapheresis is comparable to fresh one as the raw material for producing non-viral vector CAR-T.

To explore the extensive applicability of cryopreserved leukapheresis, apart from manufacturing CAR-T with non-viral vectors, we also used classical lentiviral vectors for CAR-T manufacturing. We compared the preparation of CAR-T using lentiviral vectors in T cells derived from cryopreserved and fresh leukapheresis and found no significant difference either in cell viability during the culture process (Fig. 4F) or in cell expansion (Fig. 4G). The study revealed that the proportions of Tn, TCM, TEM, and Teff (Fig. 4H) had no statistical difference between the cryopreserved and fresh leukapheresis. Certainly, when transfected at the same MOI = 1.5, there was also no significant difference in CD3 + CAR + between the cryopreserved and fresh raw materials (Fig. 4H). At the end-point of the culture, regarding the exhaustion phenotype analysis of PD1+, LAG3+, and TIM3 + cells, there was no statistical difference between the cryopreserved and fresh leukapheresis (Fig. 4I). In summary, cryopreserved leukapheresis is also on par with fresh as the raw material for producing lentiviral vector CAR-T.

To explore whether cryopreserved leukapheresis could be used as the initial raw material in the fast CAR-T process, which has more strict requirements for raw material. Our findings revealed no statistically significant differences between cryopreserved and fresh leukapheresis in terms of cell viability on the harvest day, even post-thaw (Fig. 4J). Similarly, the recovery rate after fast CAR-T preparation using the non-viral vector showed no significant difference between the cryopreserved and fresh ones (Fig. 4K). We also compared the impact of fast CAR-T preparation on cell phenotype and cytotoxicity. Post-thaw, there were no statistically significant differences in CD4 + T cell and CD8 + T cell phenotypes (Fig. 4L). Additionally, the proportions of Tn, TCM, TEM, and Teff cells were comparable, as well as between cryopreserved and fresh CAR-T (Fig. 4M). Furthermore, an assay of cellular cytotoxicity with an effector: target (E: T) ratio of 1:2 to 4:1 demonstrated no statistically significant difference in cytotoxicity efficacy between cryopreserved and fresh groups (Fig. 4N).

Cryopreserved leukapheresis as a functionally equivalent alternative to cryopreserved PBMCs for scalable immune cell therapy manufacturing

Our study compared the functional performance of cryopreserved leukapheresis and cryopreserved PBMCs as raw materials for immune cell therapies, focusing on T cells, NK cells, and NKT cells. No significant differences were observed between the two cryopreserved cell sources in multiple assays. Specifically, the proportions of CD4 + and CD8 + T cells within the CD3 + T cell population remained comparable (Fig. 5A), as did the distribution of T cell phenotypes in Tn, TCM, TEM, and Teff (Fig. 5B).

To assess functional viability, sorted T cells from both sources were cultured for 9 days. Cell viability exceeded 85% in all cases (Fig. 5C), and no differences in total T cell expansion were detected (Fig. 5D), indicating equivalent proliferative capacity. For NK/NKT cell evaluation, we employed a cytokine-based kit for NK cell expansion. However, this method concurrently amplified both NK and NKT cells, necessitating subsequent phenotypic analysis to distinguish these populations. Despite this co-expansion, viability (Fig. 5E) and total cellular expansion (Fig. 5F) during culture showed no significant divergence between cryopreserved leukapheresis and PBMCs. Further analysis of NK/NKT-specific expansion (Fig. 5G) and their proportional representation in culture up to Day 11 (Fig. 5H) also revealed comparable outcomes between the two cell sources.

These findings align with prior studies demonstrating that cryopreserved PBMCs and leukapheresis products retain functional equivalence in CAR-T manufacturing and immune cell applications, with no adverse impact on viability, expansion, or therapeutic efficacy. The observed co-amplification of NK and NKT cells highlights the importance of method-specific validation, as cytokine-driven protocols may inadvertently activate overlapping subsets. Overall, our data support the interchangeable use of cryopreserved leukapheresis and PBMCs for scalable cell therapy production.

Traditional CAR-T manufacturing uses a linear “patient-leukapheresis-immediate processing” workflow, creating geographically constrained supply chains vulnerable to cellular viability loss. Mature cryopreserved leukapheresis technology enhances flexibility by enabling centralized cryostorage combined with off-site manufacturing. Our study shows thawed products maintain sufficient viable cells (90% recovery rate, 90–99% T-cell purity) for CAR-T production. This centralized model involves cryopreserving patient materials for shipment to core facilities, genetic modification, and redistribution to clinics—distributing costs and accommodating demand fluctuations¹⁵. It facilitates equitable access for remote patients while mitigating disease progression risks associated with manufacturing delays¹⁶.

To our knowledge, this study demonstrates that cryopreserved leukapheresis products are compatible with lentiviral transduction, transposon-based systems, and rapid CAR-T manufacturing platforms, underscoring their adaptability for advancing next-generation immunotherapies. The long-term dynamics post-cryopreservation—such as in vivo expansion kinetics and the dynamics of memory subset proportions—require further elucidation through prospective clinical trials. Nevertheless, emerging clinical evidence supports the functional integrity of CAR-T products from frozen immune cells. In a multi-center trial of 115 patients receiving PBMC-derived CAR-T therapy, comparable objective response rates were observed between fresh (72%) and cryopreserved (80%) cohorts, with no statistically significant difference (p = 0.727)¹⁷. This suggests that cryopreservation protocols may preserve therapeutic efficacy. Cryopreserved leukapheresis advantages extend beyond supply chain optimization to broader clinical applications through compatibility with diverse CAR-T platforms. Studies have validated adaptability to lentiviral transduction¹ (e.g., Kymriah), transposon systems (e.g., Sleeping Beauty), and non-viral gene-editing platforms (e.g., CRISPR/Cas9)¹⁸. Cryopreserved cells show comparable therapeutic efficacy to fresh cells—demonstrated in B-cell leukemia trials with similar in vivo expansion, persistence, toxicity profiles, and disease response rates to fresh cells¹⁹. Our data reveal no significant differences in lentiviral transduction efficiency between cryopreserved leukapheresis-derived and PBMC-derived T cells. Cryopreserved umbilical cord blood-derived T cells displayed tumor-killing activity equivalent to autologous cells in preclinical models²⁰, supporting universal CAR-T development. These findings establish the cross-platform applicability, overcoming historical limitations of single-process studies.

Ultra-short cycle CAR-T manufacturing (< 3 days), exemplified by platforms like Ultra CAR-T²¹represents an emerging industry trend. Cryopreserved leukapheresis exhibits particular synergy with such technologies. Conventional ex vivo expansion (7–14 days²² risks cryopreservation-induced metabolic and phenotypic T-cell alterations, whereas fast processes mitigate these effects by minimizing culture duration (e.g., completing transduction/activation within 72 h)²³. Integration of non-viral technologies with automated closed systems (e.g., CliniMACS Prodigy) enables sub-3-day production cycles while leveraging cryopreserved leukapheresis samples for on-demand manufacturing. This “Freeze-and-Speed” strategy not only reduces cellular exhaustion risks through rapid thaw-to-manufacture workflows²⁴ but also enhances batch consistency by minimizing manual handling, thereby laying the groundwork for scalable production.

Standardized protocols for cryopreserved leukapheresis (e.g., automated apheresis devices and GMP-compliant cryopreservation) provide technical foundations for large-scale biobanking and mitigate patient-specific limitations. Critically, clinical heterogeneity is addressed through: (1) Industrial validation: Utilizing cryopreserved leukapheresis for CAR-T manufacturing achieves 82.2% release success rate across production batches for patient infusion²⁵. The Kymriah^® (tisagenlecleucel) manufacturing protocol accepts cells cryopreserved within 18 months under quality oversight²⁶; (2) Proactive banking at diagnosis: In DLBCL patients, leukapheresis prior to ASCT therapy yielded significantly increased CD4+/CD8 + ratios (p = 0.04) and reduced absolute counts of CD8 + terminally differentiated T cells (p = 0.05)²⁷, preserving T-cell fitness. This strategy enables cross-population applicability (early/late-stage patients and healthy donors), where healthy donor banking serves future universal CAR-T development. Mechanistic optimization (e.g., cryoprotectant adjustments for memory Treg subsets²⁸) will advance personalized strategies, steering CAR-T therapy from a “one-size-fits-all” toward precision medicine.

Cryopreserved leukapheresis has emerged as a cornerstone for CAR-T therapy scalability and accessibility by resolving supply chain bottlenecks, enabling cross-platform compatibility, synergizing with fast processes, and facilitating biobanking. Future research must address cryopreservation’s epigenetic impacts on immune cells and leverage artificial intelligence to optimize the entire cryopreservation-manufacturing pipeline, ultimately realizing the vision of “one leukapheresis, multiple therapies.”

Isolation and cryopreservation of PBMCs

PBMCs were isolated from leukopaks obtained from healthy donors (Milestone Biotechnologies, CN) using density gradient centrifugation. Briefly, leukapheresis samples were diluted 1:3 with normal saline and carefully layered over Ficoll-Paque Plus (Cytiva, 17-5442-03, USA). Centrifugation was performed at 800 × g for 20 min to separate PBMCs from other blood components. The PBMC layer was aspirated, washed three times with PBS, and resuspended in a freezing medium. For cryopreservation, PBMCs were adjusted to a concentration of 2 × 10⁷ – 5 × 10⁷ cells/ml in 2 ml cryovials using a cryoprotectant solution CS10 (STEMCELL, 07930, CA). The cryovials were placed in a controlled-rate freezing container and gradually cooled to − 80 °C for 24 h before long-term storage in vapor-phase liquid nitrogen tanks (Thermo). The study was conducted in accordance with the Declaration of Helsinki and approved by the Ethics Committee of Shanghai Li Quan Hospital (Continuous Review Number: 20212002-C221). Informed consent was obtained from all individual participants included in the study.

Culture of T and NK cells

T cells were magnetically sorted from PBMCs or leukapheresis products using the CliniMACS CD4/CD8 dual-positive selection system. Specifically, 2 µL of CliniMACS CD4 (Miltenyi, 200-070-213, DE) and CD8 (Miltenyi, 200-070-215, DE) microbeads were added per 1 × 10⁷ cells for positive selection, with purity validated by flow cytometry. The sorted T cells were cultured in AIM-V medium (Thermo, 0870112DK, USA) supplemented with 5% human serum substitute (Thermo, A2596101, USA), recombinant human IL-7 (25 ng/ml) (PrimeGene, GMP-101-07, CN), and IL-15 (25 ng/ml) (PrimeGene, GMP-101-015, CN). Fresh cytokines were replenished every 48 h, and cells were passaged at a 5 × 10⁵ cells/ml concentration. NK cells from PBMCs or Leukapheresis were cultured in NK expansion media (ExCell Bio, NE000-N032, CN) and as instructed.

Tumor cell line culture

The SKOV3 cell line (ATCC HTB-77) was routinely maintained in DMEM high-glucose medium (Corning, 10-013-CV, USA) containing 10% fetal bovine serum (Gibco, 10099–141 C, USA) at 37 °C and 5% CO2. Cells were passaged twice weekly using 0.25% trypsin.

CAR-T production by different techniques

T cell purification: T cells were isolated from leukapheresis-derived cells (1 × 10⁷ cells) using CliniMACS CD4 and CD8 Microbeads (2 µL each) for magnetic separation, enriching CD4⁺ and CD8⁺ populations.

T cell activation: Purified T cells were activated with CD3/CD28 TransAct (Miltenyi, 200-076-202, DE) at a concentration of 2.86 × 10⁶ cells/ml. For lentiviral vectors (LV) and PiggyBac transposon (PB) CAR-T methods, activation lasted 48 h, while fast CAR-T utilized a shorter 24-hour activation period.

Genetic modification

LV CAR-T: On day 3, T cells were transduced with lentiviral vectors (MOI 1.5) (Hillgene, HG-CT1901, CN) followed by a 9-day expansion phase before harvest.

PB CAR-T: On day 3, activated T cells were electroporated (MaxCyte GTx system) with the PB transposon vector and transposase mRNA, then cultured for 9 days.

Fast CAR-T: After 24-hour activation, T cells were electroporated in Opti-MEM using the JL transposon system and harvested immediately without further expansion.

Cell culture and harvest: LV and PB CAR-T cells underwent prolonged culture (9 days) to achieve expansion, whereas fast CAR-T bypassed amplification, emphasizing rapid production.

Cytotoxic assay

Cell toxicity was estimated by a 24-hour co-culture assay. Cell death was detected using the Real Time Cellular Analysis (ACEA XCELLIgene RTCA TP, Agilent, USA). T cells and target cells were co-cultured at an indicated E: T ratio. The day before the CAR T cytotoxicity assay, SKOV3 in good condition was inoculated into the E-Plate and set up according to the instrument instructions. The target cells were cultured overnight to achieve a cell index ≥ 1.0. Positive CAR T cells were added to the target cell culture wells for co-culture, and the experiment was completed after 24 h. Duplicate wells were performed for all samples. The percentage of the target cells killed was measured by the following method(::=frac{text{t}text{a}text{r}text{g}text{e}text{t}:text{c}text{e}text{l}text{l}:text{i}text{n}text{d}text{e}text{x}-text{t}text{e}text{s}text{t}:text{c}text{e}text{l}text{l}:text{i}text{n}text{d}text{e}text{x}}{text{t}text{a}text{r}text{g}text{e}text{t}:text{c}text{e}text{l}text{l}:text{i}text{n}text{d}text{e}text{x}}times:100).

Flow cytometry

The collection of in vitro cultured cells or isolated from leukocytes was washed twice with PBS, 1 × 10⁶ cells were resuspended in 100µL PBS, and the corresponding antibodies were added and incubated for 15 min at 4 °C. Details of the antibodies used are given in Table 3. The lymphocyte subsets were analyzed: T (CD3+), B (CD3-CD19+), NK (CD3-CD56 + CD16+), and NKT (CD3 + CD56+). T cell differentiation phenotypes were analyzed: naïve T cell (CD3 + CD45RO-CCR7+, Tn); central memory T cell (CD3 + CD45RO + CCR7+, TCM); effector memory T cell (CD3 + CD45RO + CCR7-, TEM); effector T cell (CD3 + CD45RO-CCR7-, Teff). CAR positivity was assayed using recombinant proteins. 1µL PE was added as a secondary antibody per 1 × 10⁶ cells and incubated at room temperature away from light for 15 min. Cell collection was performed using a flow cytometer (CytoFlex, Beckman, CA), and data analysis was performed using Kaluza Analysis.

Statistical analysis

The statistical analyses were performed using GraphPad Prism 8.0. Paired t-tests were used to compare one condition, while two-way ANOVA was used for comparisons involving more conditions. Error bars represent the standard errors of the mean (SEM). Statistically significant differences were indicated as follows: p ≥ 0.05 (ns, no significant difference), p < 0.05 (*), p < 0.01 (**), p < 0.001 (***), and p < 0.0001 (****).

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Author notes

1. Menglin Ren, Xiaodie Wang and Jiangrui Chen contributed equally to this work.

Authors and Affiliations

1. Shanghai Mengchao Cancer Hospital, Shanghai University, 118 Qianyang Road, Shanghai, 201805, China

   Menglin Ren, Yan Sun, Tao Liu, Qijun Qian & Zenghui Xu

2. Shanghai Cell Therapy Group Co., Ltd, Shanghai, 201805, China

   Menglin Ren, Jiangrui Chen, Yuanjian Xu, Yan Sun, Tao Liu, Qijun Qian & Zenghui Xu

3. School of Medicine, Shanghai University, Shanghai, 200444, China

   Xiaodie Wang & Qijun Qian

4. Shanghai Cell Therapy Research Institute, Shanghai, 201805, China

   Yan Sun, Tao Liu, Qijun Qian & Zenghui Xu

Contributions

M.R. wrote the manuscript. M.R., X.W., and J.C. designed and performed most of the experiments and data analysis. Y.X. assisted in project management and conducted the investigation. Y.S. and T.L. engaged in project administration and funding acquisition. Q.Q. and Z.X. provided funds, guidance, and conceptualized the research project. All authors have reviewed and approved the definitive version of this manuscript before submission.

Corresponding authors

Correspondence to Qijun Qian or Zenghui Xu.

Competing interests

The authors declare no competing interests.

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