CAR T cells with dual targeting of CD19 and CD22 in adult patients with recurrent or refractory B ce

Despite impressive progress, more than 50% of patients treated with CD19-targeting chimeric antigen receptor T cells (CAR19) experience progressive disease. Ten of 16 patients with large B cell lymphoma (LBCL) with progressive disease after CAR19 treatment had absent or low CD19. Lower surface CD19 density pretreatment was associated with progressive disease. To prevent relapse with CD19− or CD19lo disease, we tested a bispecific CAR targeting CD19 and/or CD22 (CD19-22.BB.z-CAR) in a phase I clinical trial (NCT03233854) of adults with relapsed/refractory B cell acute lymphoblastic leukemia (B-ALL) and LBCL. The primary end points were manufacturing feasibility and safety with a secondary efficacy end point. Primary end points were met; 97% of products met protocol-specified dose and no dose-limiting toxicities occurred during dose escalation. In B-ALL (n = 17), 100% of patients responded with 88% minimal residual disease-negative complete remission (CR); in LBCL (n = 21), 62% of patients responded with 29% CR. Relapses were CD19−/lo in 50% (5 out of 10) of patients with B-ALL and 29% (4 out of 14) of patients with LBCL but were not associated with CD22−/lo disease. CD19/22-CAR products demonstrated reduced cytokine production when stimulated with CD22 versus CD19. Our results further implicate antigen loss as a major cause of CAR T cell resistance, highlight the challenge of engineering multi-specific CAR T cells with equivalent potency across targets and identify cytokine production as an important quality indicator for CAR T cell potency.

Impressive antitumor effects of chimeric antigen receptor-modified T cells1,2,3,4,5,6,7,8,9,10and natural killer (NK) cells11 targeting CD19 (CAR19) have driven a paradigm shift in the treatment of relapsed or chemotherapy-refractory (relapsed/refractory) B cell malignancies. However, most patients treated with CAR19 experience disease progression. Clinical factors such as preCAR disease burden and serum lactate dehydrogenase (LDH)5,8have been linked to response to CAR19 therapies. Disease progression associated with loss of cell surface CD19 has been reported in 30–95% of relapses after CAR19 therapy in B-ALL12, through a variety of mechanisms including splice mutations and retained intracellular CD1912,13,14,15,16. Several reports have also demonstrated that effective CAR T cell responses require high target antigen expression density17,18,19,20,21; resistance associated with diminished antigen density has been shown after treatment with a monospecific CD22-CAR22 and B cell maturation antigen CARs23. While CD19 is expressed at variable levels in LBCL21, the role of emergence of CD19−/lo LBCL in CAR19 resistance has not been well studied24,25. Engineering next-generation therapeutics to overcome newly defined mechanisms of resistance is an important unmet goal.

CD22 is a sialic acid-binding adhesion molecule largely restricted to the B cell lineage and expressed on most B-lineage malignancies26,27,28,29,30. In a study of children and young adults with B-ALL enriched for patients with progressive disease after CD19-directed therapy, CD22 CAR T cells induced a 73% CR rate with equal effectiveness in CD19+ and CD19− B-ALL22,31. However, relapse was associated with the emergence of CD22lo B-ALL, illustrating the limitation of sequential targeting in B-ALL. In a phase I trial of adults with relapsed/refractory LBCL and B-ALL, we tested a bispecific CAR T targeting CD19 and/or CD22 (CD19-22.BB.z)22 and demonstrated the feasibility of manufacturing in a closed system, safety of bispecific CAR T cells and clinical activity in both B-ALL and LBCL. The observed relapses with CD19−/lo and maintained CD22 expression could be due to reduced potency of the bispecific receptor toward CD22 versus CD19.

Axicabtagene ciloleucel resistance is associated with CD19− and CD19loLBCL

To assess whether resistance to CAR19 therapy in LBCL is associated with CD19−/lo relapse, we used the semiquantitative immunohistochemistry (IHC) H-score to measure CD19 expression at baseline and at disease progression in patients treated with axicabtagene ciloleucel (axi-cel) at our institution. Our cohort comprised 44 consecutive patients treated with axi-cel with available pretreatment tissue biopsies for IHC. With a median follow-up of 21 months (95% confidence interval (CI) 11–24), median progression-free survival (PFS) was 6.1 months (95% CI 3.1–not estimable); 23 patients (52%) experienced progression. Sixteen of 23 patients with progression had postprogression biopsies available. Before axi-cel, the median CD19 H-score was 285 (interquartile range (IQR) = 240–285) (Fig. 1a and Supplementary Fig. 1). Using an H-score cutoff of 150 to indicate CD19 positivity, 39 (89%) patients were CD19+ pretherapy. CD19 positivity was not associated with durable response (Fisher’s exact test P = 1). In contrast, only 6 of 16 samples studied (37.5%) were CD19+ at disease progression. Among patients with paired pre- and posttherapy H-scores, 9 of 15 (60%) converted from CD19+ pretherapy to CD19− at relapse (McNemar test P = 0.003) (Fig. 1b). Some postprogression biopsies showed complete loss of CD19, while others demonstrated diminished CD19 expression (Fig. 1c). Additionally, treating the pretreatment H-score as a continuous variable also demonstrated no difference between patients with durable disease control versus those who experienced disease progression (t-test P = 0.32). These data demonstrate that progression after axi-cel therapy for LBCL is associated with emergent CD19−/lo disease in a high percentage of patients, but pretreatment semiquantitative IHC measurement of CD19 expression does not identify patients at risk of relapse.

To evaluate whether quantitative assessment of CD19 cell surface density might predict outcomes after axi-cel and attempt to define a threshold level of antigen expression associated with relapse, we used quantitative flow cytometry to measure CD19 site density on viable single-cell suspensions obtained by fine needle aspiration. Preaxi-cel LBCL demonstrated substantial intra- and interpatient variability in median CD19 site density as illustrated in Fig. 1d, which shows LBCLs from two representative patients, both of whom were CD19+ by IHC H-score. Patient A116 had a median CD19 site density of 6,538 molecules per cell and experienced durable disease control; in contrast, patient A140 had a median CD19 site density of 952 molecules per cell and experienced disease progression at 3 months after initial CR. Median preaxi-cel CD19 site density (n = 15) using quantitative flow cytometry was 6,396 molecules per cell (IQR = 3647–8540) with a range of 952–46,805 molecules per cell (Fig. 1e). Using a penalized logistic regression model, patients with lower pretherapy median CD19 site density had significantly increased risk of progression after axi-cel (P = 0.03), with a 50% risk of progression for patients with 2,934 CD19 molecules per cell. Therefore, we selected 3,000 molecules per cell as a cutoff to define CD19 positivity (Extended Data Fig. 1). In our cohort, 3 of 4 patients with LBCL expressing ≤3,000 CD19 molecules per cell experienced progression while 1 of 11 patients with LBCL expressing >3,000 molecules per cell had disease progression. Of 8 patients studied after axi-cel progression, 4 demonstrated a CD19 site density ≤3,000 molecules per cell (Fig. 1f and Extended Data Fig. 2). The H-score did not correlate well with antigen site density (Spearman r = 0.28, P = 0.38; Fig. 1g and Extended Data Fig. 2). These data corroborate the emergence of CD19−/lo LBCL as a major cause of resistance to CAR19 therapy, suggesting that the risk of relapse increases in LBCLs expressing a median of ≤3,000 molecules per cell before therapy, raising the prospect that quantitative flow cytometry, but not IHC, may identify LBCL patients a priori at risk of CD19−/lo relapse after CAR19.

CAR construct and clinical trial design

Previous data demonstrated that CD19 loss is an important mechanism of CAR19 failure in B-ALL. The data presented in this study similarly implicate absent or decreased cell surface CD19 as a mechanism of resistance after CAR19 in LBCL, providing a rationale for dual antigen targeting in B-ALL and LBCL. As reported previously22,32, we generated a CD19-22.BB.z-CAR comprising a single cistron encoding the anti-CD19 murine FMC63 single-chain variable fragment (scFv) and fully human anti-CD22 m971 scFv (ɑCD19 vH-ɑCD22 vL-linker-ɑCD22 vH-ɑCD19 vL), followed by human CD8 hinge and transmembrane domains, 4-1BB costimulation and CD3ζ activation domains. The CD19-22.BB.z-CAR was encoded by a self-inactivating lentiviral vector under control of an murine stem cell virus internal promoter (Fig. 2a and Supplementary Fig. 2). We conducted a phase I clinical trial of CD19-22.BB.z-CAR in patients with relapsed/refractory B-ALL and LBCL, evaluating the feasibility of manufacture and safety of CD19-22.BB.z-CAR as the primary end points. Eligible patients had disease relapsed or refractory after two or more lines of therapy and had measurable disease that expressed CD19. Patients received conditioning chemotherapy followed by CAR T cell infusion at 2 dose levels: 1 × 106 CAR+ cells kg−1 (DL1) and 3 × 106 CAR+ cells kg−1 (DL2) using a 3 + 3 dose escalation phase that enrolled LBCL and B-ALL in one cohort. While a maximal tolerated dose was not identified, a third dose level at 1 × 107 cells kg−1 was not pursued due to the efficacy seen at DL2 and concern regarding the possible toxicity of cell doses higher than 3 × 106 cells kg−1 seen in other clinical trials. Therefore, the two disease-specific expansion cohorts received the recommended phase 2 dose of 3 × 106 CAR T cells kg−1 (DL2).

a, CD19-22-CD8.BB.z-CAR contained the CD19 FMC63 and CD22 M971 scFvs, CD8α hinge and transmembrane domains, a 4-1BB costimulatory domain and a CD3ζ domain. The unique bispecific structure shows FMC63 heavy chain proximal, followed by M971 light chain, M971 heavy chain and FMC63 light chain distal. b, CAR T manufacturing and clinical trial schema. The manufacturing schema shows the TransAct process change from old to new matrix. The clinical trial schema shows the screening, lymphodepletion, CAR T cell infusion and disease evaluation time points. LP, lumbar puncture. c, Improved culture expansion resulting from the new matrix manufacturing process compared to the old matrix (P < 0.0001, two-tailed t-test). d, Significant reduction in doubling time with the new matrix process compared to the old matrix (P = 0.0411, two-tailed t-test). e, No significant difference in transduction efficiency between old and new matrix (P = not significant (NS), two-tailed t-test). Overall average transduction efficiency was 60.1% (n = 39 individual products). f, Composition of apheresis, CD4/8-enriched and CD19-22.BB.z product over time, looking at T cell (CD3+CD56−), B cell or leukemic cell (CD20+), CD4+, CD8+, NKT-like (CD3+CD56+CD16+), NK, monocyte and neutrophil subsets. g, Phenotyping of CAR T cell product reveals a skewing toward CD4+ cells (n = 39 individual products). h, Comparing the fold increase from apheresis to enrichment to final product reveals the skewing toward CD4+ cells that occurred during culture between enrichment and final product (P < 0.0001, two-tailed t-test). i, Phenotyping of T cell memory subsets revealed an enrichment in TSCM (P < 0.0001, two-tailed t-test) and TCM (P < 0.0001, two-tailed t-test) cell subsets and a depletion of the TEMRA (P < 0.0001, two-tailed t-test) subset. There was no significant change in TN or TEM cell subsets between enrichment and CD19-22.BB.z product.

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Patient characteristics

Thirty-nine patients were enrolled in two cohorts (B-ALL, n = 17, LBCL, n = 22); 38 patients received the CD19-22.BB.z-CAR infusion while one patient with LBCL died during lymphodepletion due to progressive disease and sepsis (Extended Data Fig. 3). The median age for the cohort with LBCL who received the infusion was 69 years (range 25–78) (Table 1); 15 had double expressor (expression of c-MYC and BCL2) LBCL, 3 had high-grade B cell lymphoma with translocation of c-MYC and BCL2 and/or BCL6 (double-hit) and 4 had previous autologous stem cell transplant. All patients with LBCL were CAR-naive. In the cohort with B-ALL, median age was 47 years (range 26–68); 71% had progressed after allogeneic hematopoietic cell transplantation (HCT), 65% had previous CD19-directed therapy (including 1 patient receiving previous CAR T) and 29% had previous CD22-directed therapy (Table 2). Sixty-three percent had previous central nervous system (CNS) involvement and 12% had active CNS disease at the time of enrollment.

Feasibility of closed-system manufacturing

A primary objective of this clinical trial was to assess the feasibility of cell production using closed-system manufacturing in the CliniMACS Prodigy (Miltenyi Biotec), defined as >80% of cell products meeting the protocol-specified cell dose. The manufacturing schema is illustrated in Fig. 2b and Extended Data Fig. 4a. Initially, a 7–9 d manufacturing process (old matrix) utilized washout of the T cell activator (TransAct) on day 5. At DL1, 57% (4 out of 7) of products met the prescribed cell dose within 7 d and 43% (3 out of 7) of products within 9 d. To reduce manufacturing time while increasing dose level, we incorporated a process change to remove the TransAct on day 3 (new matrix), significantly shortening the culture time required to meet the dose, which is reflected by the increased total cell count by day 7 (P < 0.0001; Fig. 2c) and decreased product doubling time (P = 0.04; Fig. 2d). Among 39 patients who underwent apheresis, CAR T products meeting predefined release criteria were successfully manufactured in 100%. Ninety-seven percent of products met the protocol-specified dose; 82% (32 out of 39) met the protocol-specified dose in 7 d (4 old matrix, 28 new matrix) (Extended Data Fig. 4b–d). Mean transduction efficiency was 60.1% (range 34.6–75.2%; Fig. 2e), with an average vector copy number of 2.23 (range 1.31–4.0; Extended Data Fig. 4e).

Characterization of CD19-22.BB.z-CAR T infusion products

Samples were obtained from apheresis, postCD4/CD8 T cell enrichment and final CAR T product collection for cell subset analysis and to confirm enrichment in T cell populations and removal of leukemic cells. Compared to apheresis products, manufactured products demonstrated increased T cell and CD4+ populations, no change in CD8+ and natural killer T (NKT)-like subsets and depletion of B or leukemic cells (defined by CD20+), NK cells, monocytes and neutrophils (Fig. 2f).

CD19-22.BB.z products showed a CD4+ predominance (Fig. 2g). To determine whether this was due to T cell enrichment postapheresis or the manufacturing process, we assessed the fold increase in CD4+ and CD8+ populations over the manufacturing process (Fig. 2h). While there was no difference in fold increase from apheresis to enrichment, there was a significant difference in fold increase between CD4+ and CD8+ T cell subsets from enrichment to final product (P < 0.0001), implicating the manufacturing process in enriching the proportion of CD4+ cells in the manufactured product. Compared to enriched apheresis samples, products demonstrated enrichment in stem cell memory T (TSCM) (P < 0.0001) and central memory T (TCM) (P < 0.0001) cell subsets, no change in naive T (TN) and effector memory T (TEM) cell populations and a decrease in terminally differentiated effector memory (TEMRA) cell subsets (P < 0.0001; Fig. 2i).

Toxicity

No dose-limiting toxicity (DLT) occurred during dose escalation and one DLT total occurred during the clinical trial. Cytokine release syndrome (CRS) of any grade occurred in 29 patients (76%) with median onset 1 d after infusion (range 0–8) and lasted a median of 4 d (range 1–12) (Tables 1 and 2 and Extended Data Fig. 5). Grade ≥3 CRS occurred in 2 patients (5%). Neurological toxicity occurred in 14 patients (37%; 9 with LBCL and 5 with B-ALL); 4 experienced ≥grade 3 neurotoxicity. Onset of neurotoxicity occurred a median of 5 d after infusion (range 3–9) and lasted a median of 4 d (range 1–11). Both CRS and neurotoxicity were treated according to institutional guidelines, with 15 patients (39%) receiving ≥1 dose of tocilizumab (range 1–3) and 45% of patients receiving corticosteroids. All episodes of CRS and neurotoxicity resolved. Two patients had laboratory evidence of macrophage activation syndrome with hyperferritinemia and hypofibrinogenemia concurrent with grade ≥3 neurotoxicity and received high-dose corticosteroids and anakinra31.

Response

The primary response assessment for LBCL was undertaken at three months after infusion. For patients with LBCL treated at the recommended phase II dose (n = 15), the overall response rate (ORR) and CR rate at 3 months were 40% (95% CI 16–68%) and 33% (95% CI 12–62%), respectively. For the entire patient cohort with LBCL (n = 21), the best ORR at any time point was 62% (95% CI 38–82%) and the CR rate was 29% (95% CI 11–52%). Five of 13 responders had improvement in response from month 1 to month 3 after infusion (Fig. 3a–c). With a median follow-up of 10 months (95% CI 8.7–21.5), median overall survival was 22.5 months (95% CI 8.3–not estimable; Fig. 3d), which could change with a longer follow-up. Median PFS was 3.2 months (95% CI 1.2–5.5; Fig. 3e). To understand the kinetics of response in patients with LBCL treated with CD19-22.BB.z, we assessed change in lymphoma burden over time by cell-free circulating tumor DNA (ctDNA) in 16 patients with available diagnostic tumor samples (Extended Data Fig. 6)33. Four patients with ongoing clinical response had no detectable ctDNA at the time of the last assessment. Among 12 patients with disease progression, we observed an initial reduction in ctDNA that nadired 14–21 d postinfusion, with 9 patients demonstrating a rise in ctDNA at or before clinical progression. These findings suggest that progressive disease after CD19-22.BB.z-CAR in LBCL is associated with a robust early response followed by early acquired resistance.

a, Swimmer plot showing the duration of remission and ongoing responses in patients with lymphoma (n = 21). Five patients had an increasing depth of response from 1 to 3 months postinfusion b, PET scans for patient S2 showing partial remission at 1 month postinfusion with subsequent progression 6 months after infusion. c, Lymphoma disease monitoring using ctDNA. After infusion of CD19-22.BB.z for patient SL02, disease burden continued to decrease; this was coincident with prolonged persistence of CD19-22.BBZ. d, Overall survival for 21 infused patients with LBCL. e, PFS for the cohort with lymphoma. f, Swimmer plot for the cohort with B-ALL (n = 17). Two patients received a consolidative allogeneic stem cell transplant (white star) g, PET scans for patient SA8, with large bulk disease preinfusion that improved to a CR 6 months postinfusion. h, Disease monitoring of patient SA8 using cellular-based NGS with sensitivity of 10−6 demonstrates increasing disease control over time coinciding with improving PET response and ongoing persistence of CD19-22.BB.z. i, Overall survival of 17 infused patients with B-ALL. j, PFS for the cohort with B-ALL. k, Absolute number of circulating CD4 and CD8 CD19-22.BB.z CAR T cells after infusion as measured by flow cytometry (n = 38 autologous infused products). l, Number of circulating CD19-22.BB.z copies per 50 ng of genomic DNA as measured by qPCR (n = 33 autologous infused products) showing initial expansion and persistence of CD19-22.BB.z as measured at 1 and 2 months (days 35–75 postinfusion), 3 months (days 76–120) and 6 months after infusion (days 120–200).

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Primary response for B-ALL was evaluated at 28 days postinfusion. All (n = 17) patients with B-ALL achieved response; 14 with CR (82%) and 3 with partial remission (Fig. 3f). One patient’s response (SA8) improved to CR 6 months after infusion (Fig. 3g,h), leading to an overall CR rate of 88%, all of whom were negative for minimal residual disease (MRD) at 10−4bone marrow sensitivity (Extended Data Fig. 7) or by positron emission tomography (PET)/computed tomography (CT) for patients with extramedullary disease. After a median follow-up of 9.3 months (95% CI 7.2–NE), median overall survival was 11.8 months (95% CI 5.5–NE) (Fig. 3i) and PFS was 5.8 months (95% CI 2.6–NE) (Fig. 3j). Two patients proceeded to allogeneic stem cell transplant in CR and are in ongoing remission. Serial MRD analyses by next-generation sequencing (NGS) of immunoglobulin receptors from bone marrow revealed that 100% (5 out of 5) of patients with ongoing CR achieved persistent MRD-negative response, while 70% (7 out of 10) of progressors had persistence or rise in MRD before or at the time of morphological relapse (Extended Data Fig. 7).

In vivo quantification of CD19-22.BB.z CAR T cells

CD19-22.BB.z-CAR cells were detected in the blood by both flow cytometry and quantitative PCR (qPCR) and peaked between days 10–14 postinfusion (Fig. 3k,l). The median peak number of circulating CD19-22.BB.z cells measured by flow cytometry was 36 CAR µl−1 (IQR = 13–136); by qPCR, it was 1,794 (IQR = 509–4,315) copies of the CD19-22.BB.z transgene per 50 ng of genomic DNA. Peak expansion did not significantly differ between LBCL and B-ALL or dose level (Extended Data Fig. 8a,b). Higher expansion as measured by area under the curve (AUC) was associated with increased CRS and neurotoxicity (Extended Data Fig. 8c,d). Despite a predominance of CD4+ cells in the manufactured CD19-22.BB.z-CAR products, CD8 CD19-22.BB.z cells demonstrated greater expansion relative to CD4 as measured by both AUC and peak levels (Extended Data Fig. 8e,f) and as seen by a peak CD4:CD8 ratio <1 in most patients (Extended Data Fig. 8g). Analysis of exhaustion markers on CAR+ cells within the product showed that CD4+ cells expressed higher levels of CD39 (ref. 34) and programmed cell death protein 1 (PD-1) (Extended Data Fig. 8h,i), potentially providing a basis for diminished in vivo expansion of CD4+ compared to CD8+ CAR T cells.

Antigen expression in patients with progressive disease after CD19-22.BB.z-CAR

We next quantified CD19 and CD22 antigen expression at the time of progression after CD19-22.BB.z-CAR. Flow cytometry, as illustrated by patient SA24 (Fig. 4a), demonstrated that 5 of 10 patients with B-ALL with progression had negative or low CD19 expression with preserved CD22, using a 90% threshold (Fig. 4b and Table 2). Paired pre- and posttherapy antigen quantification in four patients with B-ALL confirmed decreased CD19 expression in three patients with no change in CD22 expression density (Fig. 4c). In 14 patients with LBCL biopsied at progression, 3 were CD19−/lo (Extended Data Fig. 9a) by IHC. CD22 expression was not required for trial enrollment and pretreatment CD22 expression was heterogenous (Extended Data Fig. 9b); three patients had CD22−/lo LBCL and two were undetermined. All 6 patients with a pretreatment CD22 H-score >150 maintained CD22 positivity. Eleven patients with B-ALL or LBCL had quantitative flow cytometry at progression; 6 patients had low CD19 with ≤3,000 molecules per cell (Fig. 4d), including 1 patient with LBCL with a CD19 H-score >150. The median CD22 expression in patients with low CD19 was approximately 6,000 molecules per cell. Overall, 4 out of 14 (29%) patients with LBCL were CD19−/lo at progression after therapy with CD19-22.BB.z-CAR (Extended Data Fig. 9a and Fig. 4d).

CD19-22.BB.z-CAR T secretes less cytokine when stimulated through the CD22 scFv

The pattern of CD19−/lo relapse with CD22 preservation suggested that T cells expressing the CD19-22.BB.z-CAR exert significant immune pressure against CD19 but not against CD22. In vitro models demonstrated that CD19-22.BB.z-CAR was active against CD19/CD22+ cell lines (Extended Data Fig. 10a). To address this, we compared the relative potency of the signal delivered via the CD19 scFv versus the CD22 scFv within the CD19-22.BB.z-CAR (Fig. 4e) using single-cell assays. Using samples from 11 manufactured cell products, we measured the intracellular cytokine secretion (ICS) of CD19-22.BB.z-CAR T cells after coculture with double-positive NALM6 (N6-CD19/22, approximately 20,000 CD19 molecules per cell and approximately 50,000 CD22 molecules per cell), NALM6 where CD22 had undergone knockout via CRISPR–Cas9 (N6-CD19, approximately 20,000 CD19 molecules per cell, 0 CD22 molecules per cell), NALM6 where CD19 had undergone knockout via CRISPR–Cas9 (N6-CD22, approximately 40,000 CD22 molecules per cell, 0 CD19 molecules per cell) or NALM6 with both CD19 and CD22 genetically deleted (N6-double knockout) (Fig. 4f). These site densities were higher than those seen in patient samples21,28,35 and have been associated with high levels of cytokine secretion in in vitro models21.

Using ICS, CD19-22.BB.z-CAR manufactured products appeared activated as measured by CD69 expression after coculture with N6-CD19 and less activated with N6-CD22 (P < 0.0001). Higher expression with N6-CD19 compared with N6-CD22 was seen with CD107 (P < 0.0001), tumor necrosis factor-α (TNFα) (P < 0.0001), interleukin-2 (IL-2) (P < 0.0001) and interferon-γ (IFN-γ) (P < 0.0001) (Fig. 4g).

Decreased PSI and ICS resulting from CD22 as opposed to CD19 stimulation suggested decreased potency of the CD22 scFV. We next compared the activity of the CD22 scFv of CD19-22.BB.z-CAR against the identical scFV in a monospecific CD22.BB.z-CAR (Fig. 4h)22,31,36. We tested good manufacturing practice (GMP)-manufactured CD22.BB.z-CAR T cell products (n = 5) from patients with relapsed/refractory LBCL or B-ALL enrolled on an ongoing clinical trial (NCT04088890) against N6-double knockout and N6-CD22. The mean fluorescence intensity of monospecific CAR on T cells was higher than that of the bispecific CAR (Extended Data Fig. 10b,c). We found that CD19-22.BB.z-CAR and CD22.BB.z-CAR T cells manifested similar levels of activation as measured by CD69 expression and IFN-γ and CD107 secretion (Fig. 4i). However, CD22.BB.z-CAR T cells demonstrated significantly higher levels of TNF-α (P = 0.0063) and IL-2 (P = 0.0002) secretion compared to CD19-22.BB.z-CAR T cells. Similarly, single-cell cytokine secretion using the IsoPlexis platform showed a higher PSI for CD22.BB.z-CAR T cells compared to CD19-22.BB.z-CAR in the presence of N6-CD22 (n = 4; Fig. 4j). Single-cell cytokine secretion was assessed with the 32-plex human panel27 after stimulation of CD4 and CD8 CAR+ cells with the N6-CD19/22, N6-double knockout, N6-CD19 or N6-CD22 N6 lines (n = 7). Overall, CD19 stimulation with N6-CD19 yielded a higher polyfunctional strength index (PSI) testing CD8CAR+ products (P = 0.04) compared to N6-CD22 stimulation (Fig. 4j). Together, these data demonstrate that both CD19 and CD22 scFvs contained within CD19/22.BB.z-CAR can signal in response to target recognition but cytokine levels induced after CD22 scFv ligation in the bispecific CAR are lower compared to CD19 scFv ligation. In contrast, CD22 scFv ligation in the monospecific CAR resulted in higher levels of cytokines than the bispecific CD22 scFV. The reduced potency of the CD22 scFV provides a mechanism for relapse with CD19−/lo disease with preserved CD22 expression in the population studied here.

CD19 loss has been frequently identified after CAR19 in B-ALL across numerous studies incorporating variable scFvs and costimulatory domains5,6,7,15,16,37,38. In LBCL, case reports have also described CD19 loss after CAR1935,39,40,41 but a systematic analysis of CD19 expression after CAR19 in LBCL has not been conducted. In this study, we evaluated CD19 expression in 44 patients with LBCL treated with axi-cel at our institution and found that pretreatment quantitative flow cytometry was more sensitive than conventional IHC in identifying lower antigen levels associated with future progression. Quantitative flow cytometry could serve as a predictive biomarker in characterizing antigen modulation as a mechanism of resistance to CAR therapeutics.

Several studies have demonstrated the efficacy of simultaneous targeting multiple antigens in preclinical models using a variety of CAR configurations to endow multi-specific antigen recognition22,42,43,44,45,46. Early clinical trial results involving tandem CAR T cells targeting CD19 and CD20 have shown promising results with low reported rates of CD19 loss at the time of progression47,48. The CD19-22.BB.z-CAR used in this trial is a single molecule consisting of two scFvs engineered in a loop orientation22, which demonstrated activity in vitro and in xenograft models32. The clinical data presented in this study demonstrated that CD19-22.BB.z-CAR T cells were clinically active in both B-ALL, with 82% achieving an MRD−CR, and in LBCL with an ORR of 62%. Toxicity was low: 5 and 11% experienced grade ≥3 CRS or neurotoxicity, respectively. Although we observed relapses with absent or low CD19 expression, a definitive determination of whether these rates are reduced compared to those observed with monospecific CAR19 therapeutics cannot be determined in this single-arm trial given the limited data available regarding antigen−/lo relapse and the wide variability in antigen loss reported across studies.

However, the antigen expression pattern observed at relapse in B-ALL was consistent with CD19-22.BB.z-CAR delivering significant immune pressure on the CD19 antigen, whereas the lack of decrease or loss of CD22 expression suggests more limited immune pressure on the CD22 target. Experience with monospecific CD22-CAR T cells clearly demonstrated resistance associated with CD22lo B-ALL, which was not observed in this trial22. In LBCL, CD22lo disease at relapse was seen; however, CD22 expression before therapy was heterogeneous and absent in some. Our data show that CD22 scFv ligation in the bispecific CAR provided less cytokine secretion compared to CD19 scFv. These results suggest cytokine production could be a clinically meaningful product quality attribute to predict CAR potency in vivo and illustrate the challenges of delivering equivalent potency across targets in the context of multi-specific chimeric antigen receptors.

At present, much of CAR engineering is empiric and multiple approaches to simultaneous targeting of CD19 and CD22 are currently under study32,49,50,51,52. Engineering an optimal monospecific CAR is dependent on numerous factors, including prevention of tonic signaling,53 optimizing hinge length,54,55 hinge/transmembrane domain21 and optimizing the distance between the target epitope and tumor cell membrane27,56,57. In human trials, numerous chimeric antigen receptors targeting CD22 have demonstrated that small alterations in structure can result in loss of activity49,58. In the context of a tandem CAR (linked scFvs), optimal engineering also requires engineering an optimal bispecific receptor59,60, which may be particularly challenging for the CD22 target. When compared to a monospecific CAR with the same scFV, the CD19-22.BB.z-CAR demonstrated decreased TNF-α and IL-2 secretion. The threshold for CAR T cell IL-2 secretion has previously been shown to be higher than that of IFN-γ20 and may better discriminate CAR efficacy. Our results suggest that engineering iterations should be guided by careful studies of single-cell CAR polyfunctionality incorporating cytokine production as a critical quality attribute.

We also observed a significant incidence of CD19+ relapse, as reported in many previous trials of CAR19 for B-ALL and LBCL61, suggesting that potency toward the CD19 target may also be insufficient in some patients. Improvements in CAR manufacturing could prevent antigen+ resistance, which likely results from T cell failure. Defined composition of CD4:CD8 CAR cells and preferential administration of T cell subsets, such as TCM62,63, have been hypothesized to mediate optimal CAR activity1. We found our manufacturing process selectively enriched for TCM and TSCM with low TEMRA but biased the final product toward CD4 predominance. Furthermore, CD4 cells produced with our process expressed higher levels of CD39 and PD-1, which are associated with exhaustion, suggesting that the manufacturing process may not generate an optimal final product composition. Due to these findings, we paused trial enrollment to modify our manufacturing process to attain a more balanced CD4:CD8 ratio, which may also allow CD4 CAR cells to maintain a less exhausted phenotype.

In summary, this work provides evidence that antigen−/lo escape is a major pathway of resistance after CAR19 therapy for LBCL and quantitative antigen density in LBCL correlates with outcomes after CAR T cell therapy. Using a bispecific CAR capable of simultaneously recognizing CD22 and CD19, we demonstrated safety and impressive clinical activity in B-ALL. The 6-month PFS in LBCL (29%, 95% CI 12–48%) in this trial was similar to tisagenlecleucel64. Therefore, the lymphoma arm was closed in this study with enrollment ongoing in patients with B-ALL. Resistance to the bispecific CAR was associated with CD19+CD22+ relapse, probably reflecting intrinsic limitations of CAR T cells, as well as CD19−/lo but CD22+ relapse, implying inadequate immune pressure on the CD22 target. Our data also illustrate the value of cytokine production as a key product quality attribute for credentialing the potency of CAR T cells. Future work is needed to optimize multi-specific targeting by CAR T cells to improve the efficacy of this class of therapeutics both in B cell malignancies and other solid and liquid cancers.