Robust Antitumor Activity and Low Cytokine Production by Novel Humanized Anti-CD19 CAR T Cells

This article is featured in Highlights of This Issue, p. 761

Chimeric antigen receptor T-cell (CAR T-cell) therapy has shown remarkable clinical response in CD19-positive hematologic malignancies (1). Despite remarkable remission rates, limitations include life-threatening toxicities, poor T-cell persistence, immunogenic reactions, tumor escape, and technological challenges inherent to complex manufacturing processes (2–6). Moreover, the prohibitive cost of commercially available CAR T-cell therapy and the additional costs of ancillary services associated with intensive-care unit utilization make CAR T-cell therapy inaccessible to patients in countries with limited resources (7–9). Altogether, the severity of the toxicity profile along with scientific and socio-economic challenges hinders the therapeutic index and accessibility of CAR T cells in a majority of the patients who could benefit from this therapy worldwide.

Many groups reported life-threatening toxicities such as cytokine release syndrome (CRS) and neurotoxicity in large numbers of patients treated with CD19 CAR T-cell therapy (4, 10). CRS is characterized by the elevated immune response of the body that appears as high fever, inflammation, hemodynamic and respiratory compromise, macrophage activation syndrome, and vascular instability. Neurotoxicity is generally manifested by symptoms of aphasia, encephalopathy, and seizures. The onset of these toxicities has been attributed to both intrinsic and extrinsic factors. Intrinsic factors such as high disease burden during the CAR T-cell infusion are positively correlated with the severity of CRS and neurotoxicity (2). Extrinsic factors such as the design of the CAR construct play a decisive role in inducing toxicity (11). One of the most accepted plausible mechanisms for induction of toxicity is a release of a plethora of cytokines by CAR T cells upon interaction with tumor cells leading to hyperimmune activation (10). Various preclinical as well as clinical studies have demonstrated the positive correlation of severe CRS and neurotoxicity with the early elevation of proinflammatory cytokines such as IFNγ, TNFα, IL2, IL6, IL8, and IL10 post CAR T-cell infusion (12–14). Although IL6 produced by activated macrophages and by endothelial cells is believed to be the major cause of toxicities, the precise mechanisms of neurotoxicity and CRS remain unclear due to unavailability of suitable experimental models (15–17). Very recently, CD19 expression is reported on the brain mural cells, and binding of anti-CD19 CAR T cells to CD19-positive mural cells might cause neurotoxicity (17). However, mouse mural cells express negligible levels of CD19 and thus limiting the utility of the murine model in assessing the in vivo neurotoxicity. Therefore, the development of CAR T cells with a more favorable toxicity profile would extend its applicability, particularly in low-resource settings.

The attributes of cytokine secretion and antitumor efficacy of CAR T cells are largely dependent upon the anatomy and design of the CAR. We and others have highlighted that each domain of the CAR construct has a regulatory role in determining the functional efficacy and safety of the CAR T-cell therapy (1, 11, 18, 19). Very recently, just a single amino acid residue mutation in the CD28 costimulatory domain of a CAR has been shown to ameliorate CAR T-cell persistence and durability of the antitumor function compared with the native CD28 costimulatory domain (20). Another study highlighted the role of CD8α hinge and transmembrane domain of the CAR in inducing feeble T-cell activation and less cytokine secretion and thereby alleviating neurotoxicity in the patients (13). These studies indicate that even a minor alteration in CAR design strongly influences the antitumor reactivity and safety of CAR T cells.

Here, we postulated that the development and further optimization of a humanized anti-CD19 CAR could improve upon the efficacy to toxicity balance. We developed a novel humanized anti-CD19 CAR (h1CAR19-8BBζ) and demonstrated that h1CAR19 scFv interacts with CD19 antigen with higher binding affinity and more flexible complex (h1CAR19 scFv-CD19 antigen) using structural modeling and molecular dynamics (MD) simulation. Functionally, ex vivo and preclinical in vivo assessment demonstrated the robust antitumor activity of h1CAR19-8BBζ T cells with two unique features compared with the murine counterpart: (i) very low levels of cytokine production, and (2) equal distribution of CD4⁺ and CD8⁺ T cells in the final product. Further, lower cytokine production had no detrimental effect on antitumor activity and proliferation ability of CAR T cells.

The modifications to the CAR design, especially in the regulatory domains, affect the molecular structure and are critical in improving the efficacy to toxicity ratio. We designed several unique humanized scFv derived from the murine FMC63 mAb by selecting the framework of a heavy chain, VH4-34, and light chain, VK1-O18, from VBASE2 database based on the best fit similarity (21). Further, amino acid residues proximal to the complementarity determining regions (CDR), which are critical for optimal binding with CD19 antigen, were identified, and four murine residues (S²⁵, I⁶⁹, K⁷⁰, and F⁷⁸) were conserved in the humanized scFv. Both the heavy and the light chains were joined by a flexible linker (G4S)_3. Two second-generation humanized anti-CD19 CAR constructs, h1CAR19-8BBζ and h2CAR19-8BBζ, comprising humanized scFv, hinge, and transmembrane domains derived from human CD8α, human 4-1BB costimulatory domain, and human CD3ζ signaling domain were synthesized (Fig. 1A; Supplementary Table S2). The only difference between the scFv of two humanized CARs is that h1CAR19-8BBζ contains CDRs of previously published humanized anti-CD19 CAR and h2CAR19-8BBζ contains CDRs from the FMC63 mAb (22, 23).

Next, we explored the impact of CAR design on the antitumor potential of h1CAR19-8BBζ and h2CAR19-8BBζ T cells and compared it with murine anti-CD19 CAR (mCAR19-8BBζ; ref. 24). CD3⁺ T cells were isolated from peripheral blood mononuclear cells (PBMCs) of healthy donors and were transduced with CAR-encoding lentivirus, and the expression of CAR on the T-cell surface was analyzed by flow cytometry (Fig. 1B). All the constructs stably expressed the CAR on the T-cell surface [percent transduction efficiency (%TE; mean ± SEM); mCAR19-8BBζ: 28.6% ± 3.8%, h1CAR19-8BBζ: 27.2% ± 2.4%, and h2CAR19-8BBζ: 30% ± 5.90%; Fig. 1C]. Further, antigen-dependent cytotoxicity of the h1CAR19-8BBζ and h2CAR19-8BBζ T cells was determined by coculture assay using CD19-expressing tumor cell lines (NALM6, Raji, and transgenic K562CD19⁺ cells). The methodology of K562CD19⁺ cell generation is described in the Supplementary Information. Wild-type K562 cell line (CD19-ve) was used as a negative control. All the CAR T cells showed remarkable antigen-dependent cytotoxicity as quantified by flow cytometry (Fig. 1D).

Further, we quantified cytokine profile, especially IFNγ, IL2, and TNFα, secreted by CAR T cells upon antigen encounter by ELISA. Surprisingly, the h1CAR19-8BBζ T cells produced several-fold lower levels of IFNγ and decreased TNFα upon tumor cell exposure compared with the other two CARs (mCAR19-8BBζ and h2CAR19-8BBζ) in a highly reproducible manner. IL2 production was comparable and varied based on interaction with the type of tumor cell lines. The h2CAR19-8BBζ T cells showed a cytokine profile similar to mCAR19-8BBζ T cells (Fig. 1E). Despite producing low levels of cytokines, the h1CAR19-8BBζ T cells exhibited comparable in vitro expansion as well as expansion kinetics to mCAR19-8BBζ T cells (Fig. 1F).

Because the therapeutic efficacy of CAR T-cell products critically depends on the subset distribution of the final product (25), we investigated the CD4⁺ and CD8⁺ T-cell distribution in the h1CAR19-8BBζ and mCAR19-8BBζ T cells products manufactured from T cells of the same donors. Interestingly, upon flow cytometry analysis of the expanded CAR T-cell product, there was equal proliferation of both CD4⁺ and CD8⁺ T cells [CD4:CD8⁺ T-cell ratio (mean ± SEM); 0.74 ± 0.035] in h1CAR19-8BBζ T cells. However, mCAR19-8BBζ T cells showed a more skewed proliferation of CD8⁺ T cells [CD4:CD8⁺ T-cell ratio (mean ± SEM); 0.52 ± 0.024, Fig. 1G]. Thus, h1CAR19-8BBζ T cells show a more equal CD4:CD8⁺ T-cell distribution compared with mCAR19-8BBζ T cells.

Next, we examined the impact of h1CAR19-8BBζ T cells on IL6 production by monocytes using a previously reported coculture assay system, an in vitro model for CRS (15, 26, 27). The h1CAR19-8BBζ T cells or mCAR19-8BBζ T cells from multiple donors (n = 9) were cocultured with CD19⁺ Raji tumor cells, and the supernatant was collected after 24 hours. These cell-free supernatants were used to stimulate the monocytes isolated from a healthy donor. The supernatant derived from h1CAR19-8BBζ T cells induced a significantly lower amount of IL6 by the monocytes compared with mCAR19-8BBζ T cells. However, there were comparable levels of IL1β production by monocytes (Fig. 2A).

In order to examine the robustness of the h1CAR19-8BBζ T cells in vitro, we performed a repeat antigen stimulus stress test. The h1CAR19-8BBζ T cells were stimulated with CD19⁺ Raji cells with repeat stimulation after 3 days without any exogenous IL2 support. The h1CAR19-8BBζ T cells showed similar proliferation and were not exhausted after repeat tumor-specific antigen encounter similar to mCAR19-8BBζ T cells (Fig. 2B). These data suggest that lower secretion of cytokines like IFNγ had no detrimental impact on antitumor cytotoxicity and proliferation of h1CAR19-8BBζ T cells.

Further, we examined the presence of various T-cell subsets and differentiation status of T cells in CAR T cells after 14 days of expansion by multiparametric flow cytometry analysis as reported previously (28, 29). The distribution of various T-cell subsets (T_SCM: CD3⁺CD62L⁺CD45RO-CCR7⁺CD95⁺; T_CM: CD3⁺CD62L⁺CD45RO⁺CCR7⁺; and T_EM: CD3⁺CD62L-CD45RO⁺CCR7⁺) and differentiation status of T cells (based on CD45RO and CCR7 expression) in both h1CAR19-8BBζ T and mCAR19-8BBζ T cells were comparable (Fig. 2C–E), advocating that humanization preserved the stemness, memory, and effector phenotypes in both the CD4⁺ and the CD8⁺ T cells.

Moreover, in order to rule out the malignant transformation of h1CAR19-8BBζ T cells in long-term expansion, h1CAR19-8BBζ T cells were injected (subcutaneous) into immunodeficient NOD/SCID mice, and tumor growth was monitored until 4 weeks. No tumor growth at the injection site was observed in mice (Supplementary Fig. S2A and S2B).

Next, we probed the impact of CAR design on binding characteristics of humanized scFv to the CD19 target antigen. The three-dimensional structures of scFvs and CD19 antigen (PDB ID: 6AL5) were obtained using the molecular modeling as described in detail in the Methods section and in Supplementary Information (Supplementary Figs. S3–S5; Supplementary Tables S3–S6). Although molecular docking study revealed that all three scFvs prefer similar binding mode with the CD19 antigen, the lowest energy docked conformation of h1CAR19 (−21510.4 kcal/mol) compared with mCAR19 (−16684.8 kcal/mol) and h2CAR19 (−15189.2 kcal/mol) scFv with CD19 antigen (Fig. 3A). The docking analysis further indicated that h1CAR19 scFv has the lowest binding energy and establishes the maximum number of bonding interactions with the CD19 antigen compared with other scFvs and CD19 complexes (Supplementary Tables S3–S6). Next, MD simulation analysis was performed as described in detail in the Supplementary Information, indicating that all the CD19-CAR scFv complexes reached their stability after 50 ns (Fig. 3B). Interestingly, h1CAR19 scFv shows higher root mean square fluctuations compared with other scFvs (Fig. 3C; Supplementary Videos S1–S3) due to the residue composition variations in the h1CAR19 scFv, which resulted in the conformational flexibility of the heavy chain and in the linker region (Fig. 3C and D). The analysis of hydrogen-bonding interactions in the MD-simulated end structure is listed in Supplementary Tables S3–S5. Furthermore, the binding energy of CD19 antigen and scFv complexes was decreased in the order of CD19-h1CAR19 scFv (−254.57 kcal/mol) > CD19-h2CAR19 scFv (−249.18 kcal/mol) > CD19-mCAR19 scFv (−188.71 kcal/mol; Table 1), indicating h1CAR19 scFv has a higher binding affinity toward the CD19 antigen compared with mCAR19. In the CD19-h1CAR19 scFv complex, electrostatic interaction energy makes favorable contributions compared with mCAR19 and h2CAR19 complexes. Overall, our molecular modeling study confirms the higher binding affinity of h1CAR19 toward the CD19 antigen.

Given the unique profile of h1CAR19-8BBζ T cells (potent in vitro antitumor activity and very low levels of cytokine production), next, we focused on examining the antitumor activity of these cells extensively in in vivo model. To validate the in vivo functional efficacy, NALM6 cells (CD19⁺ acute lymphoblastic leukemia) bearing immunodeficient NOD/SCID mice were either treated with h1CAR19-8BBζ T cells or mCAR19-8BBζ T cells or with untransduced (UT) T cells (Fig. 4A). Tumor burden was measured weekly after tumor engraftment by bioluminescence imaging. Mice treated with h1CAR19-8BBζ T cells or mCAR19-8BBζ T cells had negligible tumor burden on day 12 compared with control groups (untreated as well as mice treated with UT cells; Fig. 4B and C). The mice treated with h1CAR19-8BBζ T cells showed significant survival benefit compared with control groups (Fig. 4D). Mice treated with repeat dose of h1CAR19-8BBζ T cells (5 × 10⁶/mouse on day 1 and day 2 after tumor cell injection) had very negligible tumor burden on day 14 compared with control groups (Fig. 4E and F). These data indicate that mice treated with h1CAR19-8BBζ T cells (either single dose or repeat dose) showed excellent efficacy and extended survival.

Further, in vivo safety profile and toxicity of h1CAR19-8BBζ T cells were determined by serum biochemistry, blood hematology, and histopathology studies of vital organs and corroborated no abnormal deviation or disease pathology in h1CAR19-8BBζ T cells–treated mice (Supplementary Fig. S2C–S2F). Upon visual inspection, the control mice depicted splenomegaly in comparison with the h1CAR19-8BBζ T-cell–treated mice group (Supplementary Fig. S2D).

Next, our goal was to assess the feasibility and efficacy of h1CAR19-8BBζ T cells generated from patients with relapsed/refractory (r/r) B-ALL in ex vivo settings. Table 3 describes the detailed characteristics of the recruited patients. In brief, patients with r/r B-ALL were defined by medullary CD19⁺ B-cell relapse after an initial remission (relapse), or by persistent bone-marrow blasts after three or more cycles of intensive chemotherapy detected by 10-color flow cytometry with minimal residual disease positivity of >0.01% (refractory). CD3⁺ T cells from the patients were isolated, activated, and transduced with h1CAR19-8BBζ construct, and their functional characterization was performed, similar to healthy donors. The transduced T cells showed the effective killing of CD19⁺ leukemia and lymphoma cells (NALM6 and Raji cells) and cytokine production (IFNγ and IL2) in an antigen-specific manner (Fig. 5A and B). No cytokine secretion and cytotoxicity were observed in CAR T cells cultured with wild-type K562 cells (CD19-ve).

Next, we optimized and validated the patient-scale CAR-T manufacturing under current good manufacturing practices (cGMP) for future clinical use. Multiple batches of h1CAR19-8BBζ T cells were manufactured in a clean environment following cGMP guidelines using a process outline in Fig. 5C. In brief, CD3⁺ T cells of healthy individuals were isolated and activated using CTS Dynabeads CD3/CD28 magnetic beads in gas-permeable cell culture bags. Next, cells were transduced with h1CAR19-8BBζ–encoding lentivirus and expanded for 9 days to achieve a clinically significant numbers. These batches of h1CAR19-8BBζ T cells showed CAR surface expression with transduction efficiency of 33.88% ± 3.26% (mean ± SEM) and consistent expansion (average 70 to 80-fold) within 9 days (Fig. 5D and E). Quality control assays for safety, identity, potency, and purity were performed for each batch to validate the process as described in Supplementary Information. Table 2 demonstrates the three representative batches of h1CAR19-8BBζ T cells qualified as per defined acceptance criteria of release testing.

CD19 CAR T-cell therapy is associated with life-threatening toxicities, which include CRS and neurotoxicity. Recent data suggest that even minor changes in CAR design can play a major role in the efficacy and safety of CARs in preclinical and clinical settings (1, 11, 13, 20, 27, 30, 31). Here, we demonstrate how our fine-tuning of the molecular structure of h1CAR19-8BBζ resulted in a favorable efficacy and toxicity profile, providing support for clinical testing of this novel construct in a low-resource setting where limited toxicity profiles will be needed to make CAR T-cell strategies more feasible.

Extensive preclinical in vivo and ex vivo data indicate that the design of the h1CAR19-8BBζ construct demonstrates the potential for a favorable efficacy. Although all three tested CAR T-cell constructs showed potent antitumor activity, there were striking differences in their structural and functional characteristics: (i) the scFv of h1CAR19-8BBζ construct formed a stable and flexible binding complex with CD19 antigen compared with the scFv of h2CAR19-8BBζ and mCAR19-8BBζ, (ii) the h1CAR19-8BBζ T cells produced significantly lower levels of cytokines (IFNγ and TNFα) compared with h2CAR19-8BBζ and mCAR19-8BBζ T cells, (iii) the monocytes produced lower levels of IL6 upon stimulation with cell-free supernatant derived from coculture of h1CAR19-8BBζ T cells with CD19⁺ tumor cells compared with mCAR19-8BBζ T cells, and (iv) the final product of h1CAR19-8BBζ T cells possesses a more even CD4⁺ and CD8⁺ T-cell distribution compared with mCAR19-8BBζ T cells.

We believe these structural and functional differences are due to the unique design of the h1CAR19-8BBζ CAR construct, which incorporates a combination of framework as well as CDRs together. Because a previous study with a humanized anti-CD19 CAR containing identical CDRs like h1CAR19 scFv showed a cytokine profile as well as a binding pattern, similar to murine mCAR19 scFv (23). Moreover, h2CAR19 scFv containing a framework identical to h1CAR19 scFv showed a binding pattern and cytokine production similar to mCAR19 scFv. However, a few studies reported that CAR scFvs with a strong affinity to tumor antigen resulted in increased “on-target off-tumor” toxicity and reduced tendency of serial killing (30, 32). Our h1CAR19 scFv forms a stable but flexible binding complex, unlike the rigid binding structure of mCAR19 and h2CAR19 scFv. These binding features of h1CAR19 scFv might be beneficial in dissociation from tumor cells and augmenting serial killing of tumor cells.

Functionally, h1CAR19-8BBζ T cells produced several folds lower IFNγ compared with h2CAR19-8BBζ as well as mCAR19-8BBζ T cells with potent in vivo and ex vivo antitumor activity. Interestingly, there were lower levels of IL6 production by monocyte stimulated with cell-free supernatants of h1CAR19-8BBζ T cells compared with murine counterparts in a coculture assay, an in vitro model of CRS (10, 15, 16). Although there are some limitations of this in vitro system due to lack of endothelial cells and other stromal elements, however, there are no suitable preclinical models to assess the CRS in in vivo settings. These data indicate that the design of the h1CAR19-8BBζ constructs favorably affected the efficacy to toxicity balance. Initially, low levels of IFNγ production by h1CAR19-8BBζ T cells were concerning as a few studies reported the positive correlation between higher cytokine production by CAR T cells with potent antitumor efficacy and positive clinical outcome (33, 34). However, similar to our observations, a very recent study showed that IFNγ-deficient CAR T cells demonstrate potent antitumor activity in a preclinical model (35). Further, recent studies demonstrated that despite low cytokine release by anti-CD19 CAR T cells and by anti-CD19 antibody TCR platform (AbTCR), these cells showed excellent antitumor activity in a clinical trial and in in vivo models respectively (13, 27). Few other studies also reported the positive correlation of increased cytokine levels especially IFNγ as one of the major factors responsible for CRS as well as neurotoxicity (10, 12, 14, 36). CAR T-cell treatment-related toxicities are a major barrier to the widespread use of CAR T-cell therapy and highlight the appropriate intervention in this direction (2, 3, 37, 38). Although various immunosuppressive agents including tocilizumab (anti-IL6R mAb), corticosteroids, and IL1 antagonists like anakinra have been shown to be effective in lowering the toxicity (10, 39, 40), the impact of these approaches on antitumor efficacy, and particularly persistence, is largely unknown and may increase the likelihood of tumor relapse and or complications from immunosuppression. Moreover, such strategies to limit toxicity may be cost-prohibitive in low-resource settings; therefore, development of a low toxicity CAR is critical.

Another interesting observation of our study was that h1CAR19-8BBζ T cells could support the proliferation of both CD4⁺ and CD8⁺ T-cell subsets depicting an equal distribution of CD4⁺ and CD8⁺ T cells in contrast to mCAR19-8BBζ T cells that supports primarily proliferation of CD8⁺ T-cell subsets. Earlier studies in preclinical as well as clinical models have established that therapeutic efficacy of CAR T-cell products critically depends on the cell phenotype, and products with defined CD4:CD8⁺ T-cell ratio demonstrate superior clinical efficacy (12, 25).

There are two additional indirect benefits of this study. First, we are developing an infrastructure for training and establishing the manpower to treat patients with CD19 CAR T-cell therapy in low-resource settings like India, where the CD19-positive malignancy burden exceeds over 30,000 patients per year (numbers are based on our experience on many large-cancer care centers). With this study, we developed a patient-scale CAR T-cell manufacturing process and established release criteria assays of identity, safety, potency, and purity required for quality control. Second, humanized anti-CD19 CAR T cells will help in reducing the immunogenicity. Clinical studies of CAR T cells with murine scFv, including Kymriah and Yescarta, reported immunogenicity as a major concern in subsets of patients (41–43). Patients treated with murine CAR T cells do not respond well to repeat infusion due to immunogenicity (12, 44, 45). However, infusion of humanized anti-CD19 CAR T cells has shown remissions in 64% of the patients who were relapsed/unresponsive to murine CAR T cells (44). In line with these studies, we believe that humanized h1CAR19-8BBζ T cells will reduce the risk of immunogenicity in case multiple dosing is required in clinical settings.

Overall, our study provides strong preclinical and ex vivo evidence for the development of a novel humanized anti-CD19 CAR with potent antitumor activity and low cytokine production that confers a favorable efficacy and toxicity profile. Several clinical trials utilizing the h1CAR19-8BBζ T cells for CD19-positive malignancies, including r/r B-ALL and diffuse large B-cell lymphoma, are planned. In addition, the capacity building (infrastructure and trained personnel) in low-resource settings like India with over 30,000 patients of CD19-positive malignancies will be beneficial in bringing CAR T-cell therapy to the majority of the patients where the toxicity of CAR T-cell therapy limits its applicability.

This study was approved by the institute ethics committee as well as by the institute biosafety committee of the Indian Institute of Technology, Bombay (IIT-B) and Tata Memorial Centre (TMC), Mumbai. The project is approved from the review committee on genetic manipulation of department of biotechnology. The blood sample from healthy donors was collected at IIT-B. The patients with r/r B-ALL (pediatric and young adults) were consented and recruited at TMC Mumbai upon obtaining informed written consent. The patient details are described in Table 3.

The humanization of two scFvs (h1CAR19 and h2CAR19) against CD19 antigen from a murine anti-CD19 scFv (FMC63 clone) was performed using the CDR grafting method (22, 46). In brief, the amino acid sequences of the FMC63 clone were subjected to Kabat numbering for the identification of CDRs. Next, the suitable acceptor humanized framework sequences were obtained using the VBASE2 database and multiple sequence alignment (21). Based on the best fit similarity with the parent antibody, the VH4-34 and VK1-O18 were identified as suitable acceptor framework sequences for the heavy chain and light chain of scFv, respectively. Further, the donor CDRs residues were grafted in the acceptor framework followed by modeling with I-TASSER server. The amino acid residues in the proximity of the CDRs were identified using Pymol software and Vernier zone identification by Kabat numbering (47). The four murine residues (S²⁵, I⁶⁹, K⁷⁰, and F⁷⁸) were identified and conserved in the humanized framework regions (Supplementary Table S2). Further, the heavy chain of scFv was linked to the light chain using a flexible (G4S)3 linker. Using this unique framework, two humanized scFvs, h1CAR19 scFv, which contains CDRs of previously published humanized CD19 CAR, and h2CAR19 scFv, which contains CDRs from the FMC63 mAb, were designed (22, 23). Next, two second-generation humanized anti-CD19 CAR constructs were synthesized; h1CAR19-8BBζ and h2CAR19-8BBζ, both consisting of hinge and transmembrane domains derived from human CD8α, human 4-1BB costimulatory domain, and human CD3ζ signaling domain, were chemically synthesized by Gene Art (Germany) and cloned in E1-T third-generation lentiviral transfer vector under EF-1α promoter.

Lentivirus production, concentration, and purification processes have been described in detail in the Supplementary Information and in Supplementary Fig. S1. Quality control assays were performed with all batches of lentivirus (Supplementary Table S1). CD3⁺ T cells were purified by EasySep Direct Human T cell Isolation Kit (Stem Cell Technologies) from the blood of the healthy donors and patients with r/r B-ALL. Purified CD3⁺ T cells were activated with CD3/CD28 dynabeads (Thermo Fisher Scientific) and transduced with CAR-encoding lentiviruses at 1,000 x g for 2 hours at 32°C on 2 consecutive days. The %TE was determined by flow cytometry using Protein-L staining. The CAR T cells were expanded for 14 days after transduction, maintaining cell density of 0.3 to 0.5 × 10⁶/mL. The phenotypic analysis of expanded cells (day 14) was performed by multiparametric flow cytometry using the staining protocol and similar gating strategies as previously reported (28, 29, 48). Data were acquired on BD FACSVerse flow cytometer and analyzed using BD FACSuite Software.

Humanized anti-CD19 CAR T cells (effector cells) were cocultured with GFP-expressing CD19⁺ lymphoma or leukemia cells as target cells (Raji, NALM6, K562CD19⁺) for 24 hours at multiple effector to target (E:T) ratios. The cell-free supernatant was collected after 24 hours for cytokines quantification, and the cell pellet was used to measure the antigen-specific killing of CD19⁺ tumor cells by flow cytometry. The cell-free supernatant was used to quantify the levels of IL2, IFNγ, and TNFα by ELISA as per the manufacturer's instructions (Thermo Fisher Scientific).

The mCAR19-8BBζ T cells and h1CAR19-8BBζ T cells generated from healthy donors (n = 9) were cocultured with Raji target cells (E:T ratio; 1:1) for 24 hours. The cell-free supernatant was collected. The monocytes were isolated from a healthy donor, and 1 × 10⁴ monocytes were stimulated with supernatant derived from cocultures of all nine donors individually for 18 to 24 hours. The cell-free supernatant from monocyte was collected and quantified for IL6 and IL1β by ELISA as per the manufacturer's instruction (Thermo Fisher Scientific).

The h1CAR19-8BBζ T cells were subjected to a repeated antigen stimulus stress test to evaluate in vitro CAR T-cell proliferation ability. The h1CAR19-8BBζ T cells were cocultured with CD19⁺ Raji cells (E:T ratio; 1:1). After an interval of 3 days, the cells were again stimulated with CD19⁺ Raji cells. At each interval, the absence of target cells was confirmed by flow cytometry, and the cells were counted using the trypan blue dye exclusion method.

To explore the binding mode of mCAR19, h1CAR19, and h2CAR19 scFv domains with CD19 antigen, we first performed molecular docking using the ClusPro 2.0 server with “Antibody mode” followed by the HADDOCK (49). This search was used to find out the interaction and orientation between the two molecules to determine the correct binding between the CD19 and scFv domains. The docked complexes of CD19 and scFv domains were further used for the MD simulation study as a starting structure. MD simulation approach (described in the Supplementary Information) was employed to investigate the refined binding mode and interaction between CD19 and scFv domains. Here, production MD simulations for 100 ns were performed for all the systems such as CD19-mCAR19-scFv, CD19-h1CAR19-scFv, and CD19-h2CAR19-scFv using the GROMACS 18.1 software. Further, analysis and visualization of MD simulation trajectories were done by using the discovery studio visualizer, chimera, and PyMolsoftware. The visual MD was used similar to the preceding study to make MD simulation movies (50).

Animal studies were performed upon approval of the institutional animal ethics committee of TMC, Mumbai. The immunocompromised NOD/SCID female mice (6–8 weeks old) were used for the study. Total body irradiation of 2.5 Gy dose was given 1 day prior to tumor injection (day -1). The following day (day 0), F-Luc⁺NALM6 cells (0.5 × 10⁶/mouse) expressing firefly luciferase fusion protein were administered by tail vein injection. The next day (day 1), mice were either treated with h1CAR19-8BBζ T cells (2.5 × 10⁶/mouse, 5 × 10⁶/mouse, or serial doses of 5 × 10⁶/mouse on days 1 and 2) or control groups [untreated controls or treated with UT cells (5 × 10⁶/mouse)] through tail vein injection. Mice treated with mCAR19-8BBζ T cells (5 × 10⁶/mouse) were used as benchmark control. The tumor burden was monitored by injecting 150 mg/kg D-luciferin (Biosynth, Switzerland) 5 to 6 minutes prior to bioluminescence imaging (Perkin Elmer IVIS 100 Imaging System). The bioluminescence flux was quantified by Live-Image software (Caliper Life Science). Mice were imaged under 2% isoflurane anesthesia and 2 L/min O₂. Mice were followed up until day 40 after tumor injection. For toxicity and safety studies, blood, bone marrow, and other vital organs such as heart, lungs, liver, lymph node, spleen, and kidneys were collected, and immunohistochemistry was performed by hematoxylin and eosin staining, and blood hematology profile and serum biochemistry were also examined.

For tumorigenicity studies, mice were injected (subcutaneous) with 5 × 10⁶ h1CAR19-8BBζ T cells and monitored for 4 weeks for any tumor development.

PBMCs were isolated from the blood of healthy donors by Ficoll-Hypaque density centrifugation method. CD3⁺ T cells were enriched and activated by CTS Dynabeads CD3/CD28 as per the manufacturer's protocol using CTS DynaMag magnet (Thermo Fisher Scientific) in CTS AIM V media (Thermo Fisher Scientific) supplemented with 5% heat-inactivated human AB serum (Valley Biomedical) containing rhIL2 (50 ng/mL, CTS Recombinant Human Protein; Thermo Fisher Scientific) at 37°C and 5% CO₂ humidified incubator for 48 hours in Permalife cell culture bags (Origen). T cells were transduced with multiplicity of infection five of lentivirus in the presence of protamine sulfate (1 μg/mL) for 2 consecutive days. The %TE was determined by flow cytometry. The CAR T cells were expanded for 9 days by maintaining the cell density of 0.3 to 0.5 × 10⁶/mL in an expansion bag (Charter Medical). On day 9, CAR T cells were counted and aliquoted for quality control assays. The methodology of quality control assays is described in Supplementary Information in detail.

Data were analyzed using GraphPad Prism version 8 and presented in the form of descriptive statistics. Error bars in each graph represent mean ± SEM. The P values were determined by unpaired t test except in Fig. 1G (paired t test). P < 0.05, 0.01, and 0.001 are represented as *, **, and ***, respectively, and P > 0.05 is considered as nonsignificant.

R. Purwar reports other from Immunoadoptive Cell Therapy Private Limited during the conduct of the study and outside the submitted work; in addition, R. Purwar has a patent for “Novel humanized anti-cd19 chimeric antigen receptor, its nucleic acid sequence, and its preparation” [patent application number: 2018 2100 5458, PCT/IN2019/050111, WO2019159193A1, WIPO (PCT)] pending and a patent for “Method for generating anti-CD19 CARs and compositions for treatment of CD19⁺ cancers using anti-CD19 CARs” (provisional patent application number: 2018 2100 5457) pending. A. Dwivedi reports a patent for “Novel humanized anti-CD19 chimeric antigen receptor, its nucleic acid sequence, and its preparation” (patent application number: 2018 2100 5458, PCT/IN2019/050111, WO2019159193A1) issued and a patent for “Method for generating anti-CD19 CARs and compositions for treatment of CD19⁺ cancers using anti-CD19 CARs” (provisional patent application number: 2018 2100 5457) pending; A. Dwivedi also holds shares in Immunoadoptive Cell Therapy Private Limited (ImmunoACT Pvt Ltd), Mumbai, Maharashtra, India. A. Karulkar reports other from Immunoadoptive Cell Therapy Private Limited outside the submitted work; in addition, A. Karulkar has a patent for “Novel humanized anti-CD19 chimeric antigen receptor, its nucleic acid sequence, and its preparation” [patent application number: 2018 2100 5458, PCT/IN2019/050111, WO2019159193A1, WIPO (PCT)] pending and a patent for “Method for generating anti-CD19 CARs and compositions for treatment of CD19⁺ cancers using anti-CD19 CARs” (provisional patent application number: 2018 2100 5457) pending. T.J. Fry reports personal fees from Sana Biotechnology outside the submitted work. No disclosures were reported by the other authors.

A. Dwivedi: Designed the experiments, interpreted the data, wrote the manuscript, designed the CAR constructs, performed in vitro assays, performed in vivo experiments, contributed to scale-up processes, and performed quality-control assays. A. Karulkar: Designed the experiments, interpreted the data, wrote the manuscript, and performed in vivo experiments. S. Ghosh: Designed the experiments, interpreted the data, wrote the manuscript, performed in vitro and in vivo assays, contributed to scale-up processes, and performed quality-control assays. S. Srinivasan: Designed the CAR constructs. B.V. Kumbhar: Performed in-silico study. A.K. Jaiswal: Wrote the manuscript, performed in vitro assays, contributed to scale-up processes, and performed quality-control assays. A. Kizhakeyil: Performed in vitro assays. S. Asija: Contributed to scale-up processes, performed quality-control assays, and wrote the manuscript. A. Rafiq: Contributed to scale-up processes and performing quality-control assays. S. Kumar: Helped in initial optimization of protocols. A. Nisar: Helped with in vivo experiments. D.P. Patil: Helped with in vivo experiments. M.V. Poojary: Obtained patient recruitment and consent. H. Jain: Obtained patient recruitment and consent. S.D. Banavali: Obtained patient recruitment and consent. S.L. Highfill: Contributed as subject matter expert, discussed the data, helped in establishing patient-scale CAR T-cell manufacturing process, and edited the manuscript. D.F. Stroncek: Contributed as subject matter expert, discussed the data, helped in establishing patient-scale CAR T-cell manufacturing process, and edited the manuscript. N.N. Shah: Contributed as subject matter expert, discussed the data, helped in establishing patient-scale CAR T-cell manufacturing process, and edited the manuscript. T.J. Fry: Contributed as subject matter expert, discussed the data, helped in establishing patient-scale CAR T-cell manufacturing process, and edited the manuscript. G. Narula: Conceptualization, supervision, edited the manuscript. R. Purwar: Conceptualization, construct design, supervision, preclinical and translational study, wrote the manuscript.

This work was funded by Tata Education and Development Trust (RD/0117TATAE00-001), Wadhwani Research Centre for Bioengineering (WRCB) at IIT Bombay (DO/2017-WRCB002-016), Tata Centre at IIT Bombay (DGDON422), and intramural funds of IIT Bombay (RD/0513-IRCCSH0-021 and RD/0115-IRSGHI0-008 to R. Purwar). Intramural grants from TMH and Tata Education and Development Trust funded the clinical studies for G. Narula. The authors would like to thank the Laboratory Animal Facility of TMC, Mumbai, for their support in conducting the animal experiments. They also thank the patients and their families, as well as the healthy volunteers who donated the samples for this study.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.