In recent years, the genomic profiling of CAR T-cell products has primarily focused on vector copy number (VCN) (9, 18, 19), and vector integration sites (8, 20, 21). Additionally, immunogenomic analyses, such as bulk and single-cell T cell receptor (TCR) sequencing (TCR-seq), have been applied to assess the TCR repertoire in CAR T-cells (22–25). For FDA-approved CAR T-cell products, VCN quantification is a mandatory component of lot-release testing, with droplet digital PCR (ddPCR) being widely used as a routine safety assay to measure VCN in most quality control (QC) laboratories (8, 9). Here, we focus on recent advancements in vector integration and TCR repertoire profiling.

On November 28, 2023, the U.S. FDA announced an investigation into cases of secondary malignancies in patients who received CAR T-cell therapy (20). The potential risk for secondary malignancies caused by insertional mutagenesis has long been a concern in CAR T-cell therapy, as viral vector transduction is required for CAR expression. However, the exact nature and frequency of genotoxicity risk associated with retroviral or lentiviral insertion remains unclear and deserves thorough investigation and transparency. Recent large-cohort follow-up studies report the incidence of secondary malignancies after CAR infusion ranging from 2-16% (26–31). Insertional mutagenesis occurs when viral vector integrates into a gene associated with cancer development, inadvertently activating an oncogene or inactivating a tumor suppressor gene and potentially leading to oncogenesis. Beyond this risk, vector integration has, in certain circumstances, been associated with a selective growth advantage, resulting in clonal CAR T-cell expansion, dominance, and persistence (32–34).

Research by Carl June’s team revealed that the integration events at the TET2 gene enhanced CAR T-cell potency (32). Similarly, an NCI research group identified clonal expansion of CAR T-cells harboring lentivector integration in the CBL gene following CAR T-cell therapy (33). Christopher et al. demonstrated that both the number and genomic loci of integration events correlate with clinical outcome in CD19 CAR T-cell products (35). Their study found that genes with integration sites enriched in responders were commonly involved in cell-signaling and chromatin modification pathways, suggesting that insertional mutagenesis in these genes promoted therapeutic T-cell proliferation. However, the consequences of viral vector integration into these reported genes have not been consistently reproducible (32). A study on the clonal dynamics of CAR T-cells over time found that not all T-cells with TET2 integration exhibit expansion, either during CAR T-cell production or after infusion (34). These findings underscore the importance of monitoring vector integration sites with potency assays in order to address safety and efficacy concerns. Our team and other research groups have developed robust pipelines for detecting viral insertion events (21, 36–40). The Bushman lab initially applied the Illumina sequencing method to investigate viral integration events in cellular products and developed the INSPIIRED pipeline (36), which enables measurement of integration events at bulk-cell resolution. Furthermore, Wenliang Wang and colleagues developed the EpiVIA pipeline, which enables detection of integration sites at the single-cell level (39). These advancements have significantly improved the feasibility of detecting integration events and facilitate their incorporation into mechanistic and safety evaluations. However, in contrast to vector copy number (VCN), which has a defined regulatory cutoff, integration site analysis currently lacks standardized criteria for determining which insertion events are definitively oncogenic and should be excluded from infusion products. Even in the case of well-characterized oncogenes such as TP53, additional mutations are often required to drive malignant transformation, as demonstrated by Perica et al. (41). As such, integration site analysis is presently better suited for informational purposes rather than serving as a standalone lot-release assay.

Another critical genomic feature is the TCR repertoire, which has been increasingly recognized as a crucial factor influencing the treatment efficacy in immunotherapies (42–45). While CAR T-cell therapy primarily relies on the target CAR expression, endogenous TCR diversity, characterized by oligoclonality and polyclonality, within the infusion products may also contribute to treatment outcomes. Recent studies have characterized the kinetic profiling of different TCR clonotypes throughout the CAR T-cell treatment process (25, 46, 47), demonstrating the cytotoxic and proliferative features of highly expanded CAR T-cell clonotypes in patients. Paired single-cell RNA analysis and TCR repertoire profiling allow for the identification of individual CAR T-cells with distinct transcriptional phenotypes (48, 49), enabling the use of TCR clonotypes as surrogate for the expansion and persistence of functional T-cell states. Qing et al. applied this paired single-cell approach to 24 infusion products and found products associated with poor clinical responses exhibited moderately reduced TCR clonotypic diversity and showed exhaustion signatures (50).

γδ T-cells present another potentially advantageous subset in the infusion products. While the majority of CAR T-cell infusion products consist of αβ T-cells, γδ CAR T-cells have demonstrated resistance to exhaustion, exhibiting lower levels of TIM3 and PD1 expression following activation (51, 52). A longitudinal analysis of CD19 CAR T-cell therapy in a chronic lymphocytic leukemia patient who achieved a durable complete response revealed the expansion of a γδ CAR T-cell population, accounting for up to 33% of all CAR⁺ cells three months post-infusion (47). Our previous work also suggests that γδ T cell in CAR T-cell products may enhance cytotoxicity and be associated with favorable clinical responses (53). Collectively, these findings highlight the importance of TCR repertoire assessment in determining the potency of CAR T-cell infusion products.

Preclinical and clinical trial data highlight the critical role of CAR T-cell differentiation states in determining therapeutic efficacy (54–56). CAR T-cell differentiation states refer to the developmental stages that T cells progress through, such as naïve, stem-cell like memory, central memory, effector memory, and terminally differentiated effector cells (57). These states are characterized by distinct gene expression profiles, functional properties, and persistence potential (58). T-cell differentiation is epigenetically programmed and maintained in progeny cells through chromatin states and DNA methylation (59). Insights from epigenomics studies have expanded our understanding of factors influencing CAR T-cell potential beyond transcriptomic profiling alone (60–70). Carlos et al. analyzed DNA methylation profiles in 114 CD19 CAR T-cell products and identified 18 distinct epigenetic loci associated with complete response (CR), event-free survival (EFS), and overall survival (OS) post-infusion. Using these CR-associated sites, they developed and validated an epigenetic signature, termed the EPICART signature, across different cohorts, demonstrating its potential as a predictor of CAR T-cell efficacy (62). Caitlin and colleagues performed a longitudinal DNA methylation assessment of CD8⁺ CD19 CAR T-cells from patients with B-cell acute lymphoblastic leukemia (B-ALL), revealing DNA methylation programs linked to a decline in CD19 CAR T-cell memory potential and the establishment of an exhaustion trajectory (63). These findings that have also been reported by others (64–66).

Epigenetic modulation has also been explored as a method to enhance CAR T-cell functionality. Yao et al. found CAR T-cells treated with low-dose decitabine (DAC, a de novo DNA methylation inhibitor) maintained higher memory-associated and lower exhaustion-associated gene expression profiles (64). Brooke and colleagues found that deleting de novo DNA methyltransferase 3 alpha (DNMT3A) in CAR T-cells prevented exhaustion and enhanced antitumor activity (65).

Beyond DNA methylation, histone modifications have been implicated in CAR T-cell function. Research has identified distinct histone markers that distinguish CD8⁺ T-cell subsets within CAR T-cell products (61). In preclinical investigations, Michel Sadelain’s group demonstrated that disrupting SUV39H1-mediated H3K9 methylation enhances the functional persistence of CD28-based CAR T-cells (67). Similarly, Mackall and colleagues restored functionality in exhausted CAR T-cells through epigenetic remodeling (68). Collectively, these findings underscore the importance of epigenomic profiling in understanding and optimizing CAR T-cell functionality. Beyond assessing product characteristics, epigenomic insights could inform potency assay development, guiding strategies to refine CAR T-cell manufacturing and enhance therapeutic efficacy. In the future, QC laboratories should consider implementing sequencing-based DNA methylation panels as potency assays, provided they can be robustly correlated with functional outputs. Alternatively, a single PCR-based assay targeting key epigenetic loci, such as those from the EPICART signature (62), could offer a more cost-effective and accessible option for routine potency testing.

Transcriptomics is a widely applied tool for analyzing gene expression (71–73), including in CAR T-cell products (5, 74–77). Both bulk and single-cell RNA sequencing (scRNA-seq) have emerged as powerful techniques for deciphering the molecular mechanisms governing CAR T-cell functionality, persistence, dysfunction, and therapeutic efficacy (50, 54, 78–82). To date, findings from transcriptomic profiling can be summarized into several key aspects.

Advancements in proteomics profiling of CAR T-cell products have been driven primarily by mass cytometry, enabling the simultaneous characterization of intracellular signaling, activation, proliferation, cytokine production, and phenotype within a single assay (81, 105–111). Several studies have focused on deciphering CAR T-cell mechanisms of action using mass cytometry.

Goldberg and colleagues developed an integrative mass cytometry panel to analyze trafficking and functional protein expression in CD19 CAR T-cells (109), identifying upregulation of activation markers (CD27, GZMB, CD69 and CD25), proliferation marker (Ki-67), and glycolysis markers (Glut1, and LDHA) in infusion products compared to baseline leukapheresis T-cells (81, 109). Salter and colleagues used mass spectrometry (MS)-based proteomics to reveal CAR T-cell activation pathways, including MAPKs (110). Hegde et al. using cytometry by time-of-flight (CyTOF) in HER-2 CAR T-cells, found lower frequencies of CD8⁺ T cells expressing PD-1 ⁺ TIM-3 ⁺, PD-1 ⁺ LAG-3 ⁺ or PD-1 ⁺ CD39 ⁺ in patients achieving CR, while higher and more variable levels were observed in those with SD and PD (108). Additionally, single-cell CyTOF analysis of day 7 circulating CAR T-cells in axi-cel-treated (a commercial CAR T-cell product) patients with large B-cell lymphoma identified three metaclusters associated with long-term clinical response (102).

Unlike previous omics approaches, metabolomics focuses on cellular metabolism, including energy metabolism (glycolysis, oxidative phosphorylation, fatty acid oxidation, etc.), mitochondrial metabolism (biogenesis, fitness, ROS production, etc.), and the analysis of other metabolites. Recently, increasing attention has been given to metabolomics profiling of CAR T-cell products, which has provided crucial insights into how metabolic fitness influences persistence, cytotoxicity, and how this correlates with overall therapeutic efficacy (80, 86, 112–115). Oxidative phosphorylation (OXPHOS) is the predominant metabolic program in memory T-cells, while aerobic glycolysis characterizes effector T-cells (116, 117). Kawalekar and colleagues demonstrated that CAR T-cells with a CD28ζ costimulatory domain primarily rely on aerobic glycolysis, whereas those with BBζ preferentially utilize fatty acid oxidation (FAO), contributing to their enhanced persistence and central memory differentiation (118). Additionally, Cappabianca and colleagues found that metabolic priming by reducing aerobic glycolysis and increasing bound NAD(P)H activity was associated with lower cytokine production, including IFN-γ, IL-2, IL-17, and TGF-β, while promoting central memory CAR T-cell expansion and persistence in GD2 CAR T-cells (119).

As for the metabolites, Paul Renauer and colleagues identified ADA and PDK1 as key metabolic regulators that enhance CAR19 T-cell cytolysis against leukemia cells (120). ADA (adenosine deaminase) catalyzes the conversion of adenosine to inosine within the purine metabolism pathway, and inosine has been reported to induce stemness features in CAR T-cells, enhancing their potency (121). Additionally, Ye and colleagues screened 27 differentially abundant metabolites in CD22 CAR T-cells with varying efficacy, identifying proline metabolism as the most significant contributor to CAR T-cell function (112). These findings suggest that detecting inosine or other metabolites in CAR T-cell supernatants could serve as a novel potency assay.

Mitochondrial properties have also been investigated in CAR T-cell products beyond energy metabolism. CAR T-cells from patients with a complete response exhibited increased mitochondrial biomass and volume compared to non-responders, correlating positively with expansion and persistence (104, 122). This enhanced persistence may be attributed to reserved bioenergetic potential, also known as spare respiratory capacity (SRC), which enables CAR T-cells to meet metabolic demand upon activation (112, 123). Additionally, mitochondrial quantities, ATP content, and the NADH/NAD ratio were found to be higher in Tscm, which are associated with superior efficacy compared to effector memory (Tem) and central memory (Tcm) T-cells (124). Overall, glycolysis, OXPHOS activity, metabolites in supernatant and mitochondrial indices could serve as potential markers for potency assay development.

As CAR T-cell therapies advance, potency assays must evolve to accurately assess the complex characteristics of these cellular products. Insights from multi- omics approaches emphasize the need for more refined potency assays that effectively capture CAR T-cells efficacy and functionality ( Figure 2 ). While further mechanistic studies are required to establish correlations between specific certain characteristics with a functional output, such as epigenomic profiles to IFN-γ secretion or cytotoxicity, it remains valuable to broadly explore CAR T-cell characteristics for their potential for potency assay development. Some of these characteristics require several days to complete which may prevent them from being used for potency assessment at this time. However, the field is progressing rapidly and if a specific profile is found to be useful for assessing CAR T-cell potency, alternative platforms that allow for more rapid testing will likely be developed. Here, we summarize key considerations for optimizing potency assays based on recent advancements.

The emergence of non-viral gene delivery methods (e.g., transposon systems, CRISPR-mediated knock-in) has provided alternative strategies for generating CAR T-cells without the need for viral vectors (140–142). These methods generally exhibit lower transduction efficiency than viral vector methods due to reduced knock-in rates, leading to a smaller proportion of CAR-expressing cells (140). In transposon-based systems (Sleeping beauty and PiggyBac), VCN varies widely due to uncontrolled transgene integration, resulting in heterogenous CAR expression across cells. Unlike viral vector-based CAR T-cells (VCN: 1–2 copies/cell), transposon-based CAR T-cells can exhibit 0-10+ copies/cell (18, 19, 130, 143, 144), raising concerns regarding product consistency and regulatory compliance. Given the FDA’s recommendation that the VCN should remain <5 copies/genome in infusion products (8), potency assays for transposon-based CAR T-cells must account for VCN thresholds to ensure safety and efficacy. For CRISPR-mediated CAR knock-in, off-target genome edits remain a significant concern, potentially affecting T-cell function and stability. Therefore, potency assays for these CAR T-cells should include whole-genome sequencing (WGS) (131, 145, 146) or GUIDE-seq (147, 148) to accurately identify and characterize these off-target sites.

In vivo CAR T-cell generation eliminates the need for ex vivo expansion, shifting potency assessment from traditional pre-infusion characterization to real-time in vivo monitoring (149–151). In this instance, the viral vector itself is the primary product, making vector characterization an essential component of potency assay assessment. Conventional potency assays, such as those measuring transduction efficiency, cytotoxicity, and cytokine secretion in vitro, are no longer applicable. Future efforts in vitro potency analysis should focus on linking the characteristics of viral vectors and their functional outputs.

Ultra-rapid CAR T-cell manufacturing (3 days or less) (138, 152–154) significantly shortens the ex vivo expansion phase, introducing unique challenges for potency assessment. With some protocols completing the process in as little as 24 hours, the limited cell yield poses a challenge for conducting potency assays. Additionally, it remains uncertain whether these cells achieve sufficient CAR expression and vector copy number to be reliably detected by flow cytometry and ddPCR. Moreover, the shortened manufacturing time may result in a higher proportion of less-differentiated and less cytotoxic T-cells (153, 155), potentially biasing cytotoxicity potency assays for these CAR T-cells.