Chimeric antigen receptor (CAR) T-cell therapies that target either CD19 or CD22 alone have potent antilymphoma effects. However, antigen escape–mediated relapse often occurs. CAR T cells targeting both CD19 and CD22 may overcome this limitation. In this study, we developed bispecific CAR T cells simultaneously recognizing CD19- and CD22-expressing targets and assessed their safety and efficacy profiles in patients with relapsed/refractory aggressive B-cell lymphoma. Twenty-four patients were screened, and 16 were found eligible for the study. CAR T-cell–associated toxicities were recorded. Responses, overall survival (OS), and progression-free survival (PFS) were assessed. Of the 16 eligible patients, 14 (87.5%) achieved objective response and 10 (62.5%) achieved complete response (CR). The 2-year OS and PFS rates were 77.3% and 40.2%, respectively. Achieving CR (P = 0.046) and the number of prior chemotherapy lines (n = 2; P = 0.047) were independent prognostic factors associated with favorable PFS. The 2-year OS and PFS among patients who achieved CR were higher than among those who did not (P = 0.015 and P < 0.001, respectively). The 2-year PFS among patients who received two prior lines of chemotherapy was higher than that among patients who received more than two lines of chemotherapy (P = 0.049); OS did not differ between the groups. Severe grade 4 cytokine-release syndrome (CRS) was observed in 1 patient; 4 and 11 patients had grades 1 and 2 CRS, respectively. No patients developed neurotoxicity. CD19/CD22 dual-targeted CAR T cells may be a safe, potent antilymphoma cell-based targeted immunotherapy.

Chimeric antigen receptor (CAR) T-cell therapy has revolutionized the treatment of relapsed/refractory (R/R) B-cell hematologic malignancies, primarily acute lymphoblastic leukemia (ALL) and B-cell non-Hodgkin lymphoma (NHL). CD19-targeted CAR T cells yield complete remission (CR) rates of about 90% in R/R ALL, but substantially lower rates in R/R NHL (50%; refs. 1–7). Antigen escape–mediated relapse is a major limitation to long-term responses. Mechanistically, CAR T cells targeted to only CD19 exert selective pressure, leading to CD19-negative malignant clones that can escape CD19-targeted CAR T cell–mediated cytotoxicity. Several studies show that among patients with B-cell NHL who receive CD19-targeted CAR T-cell therapy, approximately 30% have CD19-negative relapse (8, 9).

In addition to the CD19-targeted CAR T cells, we developed highly active human-derived CAR T cells targeting CD22 (2). These cells yield a CR rate of 80% in patients with R/R ALL who have insufficient responses to CD19-targeted CAR T cells. The CR rate achieved with CD22-targeted CAR T cells is comparable with that with CD19-targeted CAR T cells. Despite this potent antitumor activity, some tumors cells show reduced surface expression of the CD22 antigen. Thus, CAR T cells targeting both CD19 and CD22, so-called CD19/CD22 dual-targeted CAR T cells, may have broad activity to overcome this limitation (10, 11).

Evidence in preclinical models of solid tumors shows that dual- or multi-antigen–targeting CAR T cells may exhibit synergistic effects, permitting optimization of response rates compared with those achieved from targeting single antigens (12, 13). B-cell lymphoma cells usually coexpress high levels of both CD19 and CD22. In addition, both CD19 and CD22 are validated targets for CAR T-cell therapy in B-cell hematologic malignancies (14, 15). Thus, dual targeting of CD19 and CD22 by CAR T cells may mitigate this baseline variability in antigen expression and changes that occur over time. In addition, simultaneous targeting of both CD19 and CD22 may reduce the likelihood of antigen-loss variants, similar to recent studies of bispecific CAR T cells targeting CD123 and CD19 (16).

In this study, we sought to develop bispecific CAR T cells that could concomitantly recognize CD19- and CD22-expressing targets by incorporating both CD19 and CD22 single-chain variables in a single CAR construct. We evaluated the safety and efficacy of these CD19/CD22 dual-targeted CAR T cells in 16 patients with R/R aggressive B-cell lymphoma.

Generation of CD19/CD22 dual-targeted CAR T cells

A lentiviral vector was used to carry a dual antigen–specific CAR targeting CD19 and CD22. The vector was arranged in a novel Boom configuration (patent pending), with a common 4-1BB costimulatory domain and a CD3ζ signaling domain (Fig. 1A). The antigen-recognition domain is composed of two single-chain fragment variable domains (scFv) for CD19 and CD22, respectively, that were derived from the FMC63 murine clone and the human phage display antibody library, respectively. The monospecific CAR vectors against CD19 and CD22 (CD19-BB-002 and CD22-BB-002, respectively) were constructed using the same antigen-recognition domain, as well as costimulatory and signaling domains with the bispecific CAR, as described previously (2, 17).

Figure 1.

CAR construct and clinical protocol. A, Schematic diagram of the Boom-configured CD19/CD22 dual-targeted CAR. B, The timeline of the study design. FC, fludarabine and cyclophosphamide.

Figure 1.

CAR construct and clinical protocol. A, Schematic diagram of the Boom-configured CD19/CD22 dual-targeted CAR. B, The timeline of the study design. FC, fludarabine and cyclophosphamide.

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Lentiviral vectors were generated by triple transfection of the recombinant vectors, psPAX2 (catalog no. 12260, Addgene) and pMD2.G (catalog no. 12259, Addgene), into HEK293T cells and concentrated by ultracentrifugation at 50,000 × g for 2 hours, at 16°C. CAR T-cell transduction and expansion were performed as described previously (2, 17). Briefly, peripheral blood mononuclear cells collected from healthy donors or patients were stimulated with magnetic beads coated with anti-CD3/CD28 (Thermo Fisher Scientific) overnight. The next day, transduction was performed at a multiplicity of infection ration of 1:10. Transduced cells were cultured in X-VIVO 15, a serum-free medium (Lonza) with 300 IU/mL IL2, for the duration of cell culture. Transduction efficiency and cell viability were examined at the time of cell infusion. Transduction efficiency was defined as the ratio of CAR T to total CD3+ T cells, as determined by flow cytometry. Cell viability was determined by Trypan blue exclusion.

Flow cytometric analysis of CD19/CD22 dual-targeted CAR T cells

T cells or CAR T cells were evaluated for CD3, CD4, CD8, CD45RA, and CD62 L expression using allophycocyanin (APC)-labeled mouse anti-human-CD3 (catalog no. #555335, BD Biosciences), APC-Cy7–labeled mouse anti-human CD4 (catalog no. 557871, BD Biosciences), PE-CF594–labeled mouse anti-human CD8 (catalog no. 562282, BD), BV421-labeled mouse anti-human CD45RA (catalog no. 562885, BD Biosciences), and BV650-labeled mouse anti-human CD62 L (catalog no. 563808, BD Biosciences), respectively. CAR expression was examined by staining with biotinylated goat anti-mouse and anti-human antibodies (Jackson ImmunoResearch Laboratories, Inc.), followed by APC-labeled streptavidin (BD Pharmingen). All flow cytometry readings were performed on the BD FACSCelesta system and analyzed using FlowJo software. B cells were evaluated for CD19 and CD22 expression using FITC-labeled mouse anti-human CD19 (catalog no. 302206, Biolegend) and PE-labeled mouse anti-human CD22 (catalog no. 302506, Biolegend).

Evaluation of functional characteristics of CD19/CD22 dual-targeted CAR T cells

The in vitro cytotoxic capabilities of the CD19/CD22 dual-targeted CAR T cells were assessed using HeLa cells engineered to express CD19, CD22, or both antigens at an effector-to-target (E:T) ratio of 1:1. Cytotoxicity was evaluated using a label-free iCELLigence real-time cell analyzer (RTCA) system (Agilent Biosciences, Inc.). HeLa-CD19, HeLa-CD22, and HeLa-CD19/CD22 cells were seeded at 1 × 104 per well in the RTCA unit and cultured for 24 hours. Control T cells or CAR T cells were then added at 2 × 104 per well at 100 μL/well in volume to the RTCA unit. Impedance signals were recorded for 48 hours at 5-minute intervals. At 24 hours after addition of control T cells or CAR T cells to the target cells, culture supernatant samples were collected by centrifugation at 400 × g for 10 minutes at 4°C, and were evaluated for the cytokine/chemokine concentrations using the Human XL Cytokine Magnetic Luminex Performance Assay 45-plex Fixed Panel (catalog no. LKTM014, R&D), according to the protocol provided by the kit. We evaluated the concentrations of 45 cytokines in the supernatants by using Luminex Assays (Thermo Fisher Scientific).

Cell lines and culture

HEK293T, HeLa cells, and Nalm 6 cells were purchased from the Cell Bank of the Chinese Academy of Sciences in 2014. HEK293T, HeLa cells were maintained in DMEM high glucose (Biological Industries) supplemented with 10% heat-inactivated FBS (Gibco). Nalm 6 cells were maintained in RPMI1640 (Biological Industries) medium supplemented with 10% heat-inactivated FBS. All cells were maintained in a humidified incubator with 5% CO2 at 37°C. Cell lines were not authenticated since purchase and were cultured for fewer than 10 passages. Cell lines were routinely tested for Mycoplasma using a mycoplasma contamination detection kit (rep-pt1, InvivoGen).

Generation of stable cell lines

Full-length human CD19 (NM_001178098.2) and CD22 (NM_001185099.2) cDNAs were first cloned into the lentiviral vector pWPXL (catalog no. 12257, Addgene). HEK293T cells were then transfected with the plasmid mixture of the lentiviral transfer and packaging vectors, psPAX2 (catalog no. 12260, Addgene) and pMD2.G (catalog no. 12259, Addgene), using polyethylenimine (MW 40,000, catalog no. 24765-2, Polysciences) following the manufacturer's instructions. Lentivirus-containing supernatants were harvested 48 to 72 hours after transfection. HeLa cells were then transduced with these lentiviruses, and CD19, CD22, or CD19/CD22 double positive cells (HeLa-CD19, HeLa-CD22, and HeLa-CD19/CD22 cells, respectively) were enriched by cell sorting using a cell sorter (S3e Cell Sorter, Bio-Rad) after staining with anti-CD19-PE (catalog no. 340364, BD Biosciences) and/or anti-CD22-PE (catalog no. 562859, BD Biosciences). Nalm6 cells stably expressing a fusion protein of eGFP (sequence from pcDNA3-eGFP, catalog no. 13031, Addgene) and firefly luciferase (XM_031473197.1) was engineered (Nalm6-eGFPLuc) by cloning the cDNAs of both genes into the lentiviral vector pWPXL. Lentiviruses were produced as described above. Nalm6 cells stably expressing eGFP and firefly luciferase (Nalm6-eGFPLuc) were obtained by lentivirus transduction, followed by cell sorting using eGFP as a fluorescent marker.

Xenograft animal model studies

For the xenograft animal model studies, 4- to 6-week-old NOD/SCID female mice (purchased from Hangzhou Ziyuan Experimental Animal Technology Ltd.) were implanted with 1 × 106 Nalm6-GFPluc cells per mouse through tail vein injection. After 6 days, every mouse was injected intravenously with 1 × 107 CD19/CD22 dual-targeted CAR T cells or unmodified T cells from healthy donors. Bioluminescent imaging was performed weekly using an IVIS Imaging System (PerkinElmer). Imaging was conducted 30 minutes after mice were injected intraperitoneally with the substrate for firefly luciferase, D-luciferin (Goldbio, catalog no. 1151-44-35-9), under isofluorane anesthesia. All animal studies were conducted under a protocol approved by the Animal Care and Use Committee of Zhejiang University.

Clinical trial protocol design and development

The clinical trial (ChiCTR1800015575) was designed to assess the safety and efficacy of autologous CD19/CD22 dual-targeted CAR T–cell therapy in 16 patients with R/R aggressive B-cell lymphoma. The inclusion criteria were as follows: (i) patients younger than 70 years of age; (ii) relapsed or refractory disease, defined as progressive or stable disease on the most recent chemotherapy regimen and disease progression or relapse within 12 months after autologous stem cell transplantation (diagnosis according to the WHO classification for tumors of the hematopoietic and lymphoid tissues); (iii) the presence of measurable disease; (iv) CD19 or CD22 expression on at least 90% of lymphoma cells on IHC assay or flow cytometry; (v) Eastern Cooperative Oncology Group performance status less than 2; (vi) total bilirubin ≤1.5 times the institutional upper limit of normal, aspartate transaminase (AST or SGOT) ≤3 times institutional upper limit of normal, alanine transaminase (ALT or SGPT) ≤3 times institutional upper limit of normal, serum creatinine ≤2 times the institutional upper limit of normal; and (vi) life expectancy of 12 weeks or more. Patients with uncontrollable infection, active graft-versus-host disease, or clinically evident neurologic lesions were excluded. Cytokine release syndrome (CRS) was assessed and graded according to American Society for Transplantation and Cellular Therapy (ASTCT) criteria (18). Neurologic toxicity and other adverse events were assessed and graded according to the NCI Common Terminology Criteria for Adverse Events (CTCAE), version 5.0. Blood samples were collected and analyzed freshly by flow cytometry. The protocol was approved by the Institutional Review Board of the First Affiliated Hospital of Zhejiang University School of Medicine. The trial was conducted in accordance with the principles of the Declaration of Helsinki. All enrolled patients provided written informed consent.

Cellular production and administration

Patients underwent a single leukapheresis procedure 7 to 10 days prior to CAR T-cell infusion (Fig. 1B). Peripheral blood mononuclear cells (PBMC) were isolated, activated, and transduced with a clinical-grade pseudotyped lentiviral vector that was produced according to current good manufacturing practices as described in Materials and Methods. CAR T cells were transduced, cultured, expanded, and detected as described in Materials and Methods (2, 17). Lymphodepletion included fludarabine 30 mg/m2 × 3 days (days −4 to −2) and cyclophosphamide 500 mg/m2 × 2 days (day –3 to −2), followed by infusion (day 0) of 1 to 10 × 106 fresh CD19/CD22 dual-targeted CAR T cells per kg body weight (Fig. 1B). Following administration of CAR T cells, the percentage of CAR T cells in the peripheral blood and the levels of serum cytokines were monitored on days 1 to 14 after CAR T-cell infusion.

Following CAR T–cell infusion, patients were followed up periodically at 1, 3, 6, 12, and 24 months or until disease progression. The primary objective of the study was to assess the safety of the bispecific CAR T-cell therapy in patients with R/R B-cell lymphoma. Hence, the incidence, severity, and grade of CRS; damage to major organs (heart, liver, and kidney); and vital signs were strictly monitored. The secondary end point was overall response rate to the treatment. Treatment responses were evaluated according to PET imaging and the Newly Revised Response Criteria for Malignant Lymphoma for the assessment of CR and partial response (PR; refs. 19, 20) at 1 and 3 months after CAR T-cell infusion.

Statistical data analysis

The primary endpoints were overall survival (OS), progression-free survival (PFS), and duration of response (DOR). The secondary endpoints were incidences of CAR T-cell therapy–associated toxicity and adverse events. All measurement data were described using medians and ranges and were compared using Kruskal–Wallis tests. Enumeration data were presented as frequencies (%) and compared using chi-square tests. The correlations were calculated by rank-based Spearman tests. Furthermore, the OS, PFS, and DOR probabilities were determined by the Kaplan–Meier method. All enrolled patients were involved in determining OS and PFS, whereas only patients achieving responses were included in calculating DOR. Cox regression and logistic regression models were used to obtain HR estimates and corresponding 95% confidence intervals (CI) for OS, PFS, DOR, and best objective response (OR). Finally, results were considered statistically significant if two-sided P values were less than 0.05. Data were analyzed using IBM SPSS Statistics 24 and GraphPad Prism 9.0.

Preclinical cytotoxicity evaluation of CD19/CD22 dual-targeted CAR T cells

Preclinical cytotoxicity evaluation of the CD19/CD22 dual-targeted CAR T cells was performed in comparison with monospecific CD19-BB-002 and CD22-BB-002 CAR-T cells in HeLa cells that were engineered to express CD19, CD22, or both antigens. The dual-antigen targeting configuration of the two scFvs did not interfere with the expression of the CAR on T cells (Fig. 2A). In an in vitro cytotoxicity assay, the dual antigen–specific CAR T cells performed equally well when compared with the monospecific CAR T cells when there was only a single antigen present on the target cells; better performance was observed when both antigens were present on the target cells (Fig. 2B). In addition, the dual antigen–specific CAR T cells induced equal amounts of IL3, GM-CSF, and IFNγ, when compared with the two monospecific CAR T cells (Fig. 2C). The monospecific CD19-targeted CAR T cells induced more IL2 and TNFα than the monospecific CD22-targeted CAR T cells and dual antigen–specific CAR T cells. However, in the presence of both CD19 and CD22 antigens, the dual antigen–specific CAR T cells tended to produce more granzyme B, which may explain the higher degree of cytotoxicity when compared with the two monospecific CAR T cells (Fig. 2C). Using a live imaging xenograft animal model, we found that CD19/22 dual-targeted CAR T cells were capable of effectively eradicating Nalm6-GFPluc cells in NOD/SCID mice (Supplementary Fig. S1).

Figure 2.

Preclinical evaluation of CD19/CD22 dual-targeted CAR T-cells and a brief outline of the whole procedure. A, Flow cytometry plots of CAR expression on human T cells after lentiviral transduction. Gates were set on the control untransduced CD3+ T cells. CAR expression level of nontransduced T cells, T cells transduced with CD19BB-002, T cells transduced with CD22BB-002, and CD19/CD22 Boom–transduced T cells are shown. Results are representative of three independent experiments. B, Cytotoxicity of CAR T cells against HeLa cells expressing CD19, CD22, or both CD19 and CD22. In vitro cytotoxicity of monospecific CD19-BB-002, monospecific CD22-BB-002, or CD19/CD22 Boom–targeted CAR T cells against target cells is shown. E:T ratio was 1:1. Cytotoxcity was assessed using the RTCA system (Agilent Biosciences Inc.). Results are representative of three independent experiments. C, Levels of the indicated cytokines and chemokines in supernatants from the cytotoxic assay. The cultured supernatant samples of CAR T cells cocultured with target cells were harvested after 24 hours and tested for 45 cytokines/chemokines. Top, the heat map of 45 cytokine/chemokine of the culture supernatant samples. Bottom, bar graphs for selected cytokines: IL2, IL3, TNFα, GM-CSF, granzyme B, and IFNγ (n = 3). This assay was done in triplicate. Error bars represent SD. One-way ANOVA with Kruskal–Wallis test was used for statistical analyses, with P value threshold <0.05. ns, not significant. *, **, ***, and **** stand for P < 0.05, P < 0.01, P < 0.001, and P < 0.0001, respectively. D, A brief outline of the whole procedure in the study.

Figure 2.

Preclinical evaluation of CD19/CD22 dual-targeted CAR T-cells and a brief outline of the whole procedure. A, Flow cytometry plots of CAR expression on human T cells after lentiviral transduction. Gates were set on the control untransduced CD3+ T cells. CAR expression level of nontransduced T cells, T cells transduced with CD19BB-002, T cells transduced with CD22BB-002, and CD19/CD22 Boom–transduced T cells are shown. Results are representative of three independent experiments. B, Cytotoxicity of CAR T cells against HeLa cells expressing CD19, CD22, or both CD19 and CD22. In vitro cytotoxicity of monospecific CD19-BB-002, monospecific CD22-BB-002, or CD19/CD22 Boom–targeted CAR T cells against target cells is shown. E:T ratio was 1:1. Cytotoxcity was assessed using the RTCA system (Agilent Biosciences Inc.). Results are representative of three independent experiments. C, Levels of the indicated cytokines and chemokines in supernatants from the cytotoxic assay. The cultured supernatant samples of CAR T cells cocultured with target cells were harvested after 24 hours and tested for 45 cytokines/chemokines. Top, the heat map of 45 cytokine/chemokine of the culture supernatant samples. Bottom, bar graphs for selected cytokines: IL2, IL3, TNFα, GM-CSF, granzyme B, and IFNγ (n = 3). This assay was done in triplicate. Error bars represent SD. One-way ANOVA with Kruskal–Wallis test was used for statistical analyses, with P value threshold <0.05. ns, not significant. *, **, ***, and **** stand for P < 0.05, P < 0.01, P < 0.001, and P < 0.0001, respectively. D, A brief outline of the whole procedure in the study.

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Patient baseline characteristics

From December 1, 2017, to September 30, 2019, 24 consecutive patients were screened. Six patients were not eligible for the clinical trial owing to respiratory failure (1 patient), severe infection (two patients), disease progression (2 patients, liver failure due to liver metastasis), and renal failure (one patient). Thus, 18 patients were enrolled in the study and underwent leukapheresis (Fig. 2D). Of these 18 patients, two patients underwent leukapheresis but subsequently withdrew from further studies owing to severe infection (one patient) and disease progression (one patient). Four patients were treated in the proof-of-concept (POC) procedure before trial registration in the Chinese Clinical Trial Registry (ClinicalTrials: ChiCTR1800015575).

Patient baseline characteristics are summarized in Supplementary Table S1. With regard to the type of NHL, 10 (62.5%) patients had nongerminal center B cell–like disease, two (12.5%) had germinal center B cell–like disease, two (12.5%) had B-cell lymphoblastic lymphoma, and one (6.25%) had Burkitt's lymphoma. All patients had advanced disease and had been heavily pretreated with various regimens, including chemoimmunotherapies such as anti-CD20 therapy, alkylating agents, and anthracycline. All patients were refractory and progressed on the last course of therapy. Thirteen were not eligible for autologous hematopoietic stem cell transplantation (HSCT) as they had refractory disease; three had experienced relapse after multiple lines of chemotherapy (one following autologous HSCT). Fifteen (93.8%) patients had stage III or IV disease, and only one patient had stage I disease. Eleven (68.8%) of the patients had extranodal infiltration. The median number of prior therapies was 3. The features of the four patients from the POC step were consistent with those of the 12 patients enrolled after trial registration in terms of inclusion/exclusion criteria, CAR T-cell manufacture protocol, study protocol, follow-up scheme, and adverse effect management principle (see the detailed clinical trial protocol on the ChiCTR website). There were no significant differences in patient characteristics between the POC and clinical trial groups (Supplementary Table S2).

Characterization of CD19/CD22 dual-targeted CAR T-cell products

The manufacturing process for the CD19/CD22 dual-targeted CAR T cells was successful for all patients. The incubation time was 7 to 11 days. The median transduction efficiency of the final products was 48.5% (22%–70%) by flow cytometry (Table 1). The median CAR T–cell infusion dose was 6.3 × 106/kg (4.9–9.4 × 106/kg). The median ratio of CD4+ to CD8+ CAR T cells in the infusion cells was 2.05 (0.67–9.57; Table 1). The median proportion of CD4+CD45RA+CD62L+ CAR T cells among CD4+ CAR T cells was 45.05% (7.1%–63.1%). The median proportion of CD8+CD45RA+CD62L+ CAR T cells among CD8+ CAR T cells was 74.5% (31.0%–94.6%; Table 1).

Table 1.

CD19/CD22 dual-targeted CAR T-cell characterization in CAR T-cell products.

Patient no.Transduction rate (%)CD4/CD8 ratio in CAR T cellsCD4+ CD45RA+ CD62L+ in CD4+ CAR T cells (%)CD4+ CD45RACD62L+ in CD4+ CAR T cells (%)CD8+ CD45RA+ CD62L+ in CD8+ CAR T cells (%)CD8+ CD45RACD62L+ in CD8+ CAR T cells (%)
22 1.8 7.1 51.7 31.0 68.4 
38 1.4 63.1 17.9 92.9 7.1 
65 9.6 23.8 24.3 54.3 44.9 
44 5.4 61.4 22.5 88.0 12.0 
46 4.0 22.8 22.0 74.4 23.5 
57 0.8 55.1 11.7 74.6 24.3 
52 0.7 43.8 24.1 80.0 19.9 
65 2.3 58.9 35.6 90.9 8.9 
57 3.7 60.0 28.9 89.6 9.6 
10 34 2.4 40.4 22.5 73.4 26.4 
11 70 0.9 46.3 24.7 66.2 32.4 
12 49 1.0 49.4 17.8 78.2 20.5 
13 30 2.3 53.5 4.2 94.6 1.6 
14 55 0.7 28.5 27.9 67.6 19.4 
15 45 3.0 9.5 5.4 71.6 26.9 
16 48 0.9 35.5 28.6 73.0 27.0 
Median 48.5 2.1 45.1 23.3 74.5 22.0 
Patient no.Transduction rate (%)CD4/CD8 ratio in CAR T cellsCD4+ CD45RA+ CD62L+ in CD4+ CAR T cells (%)CD4+ CD45RACD62L+ in CD4+ CAR T cells (%)CD8+ CD45RA+ CD62L+ in CD8+ CAR T cells (%)CD8+ CD45RACD62L+ in CD8+ CAR T cells (%)
22 1.8 7.1 51.7 31.0 68.4 
38 1.4 63.1 17.9 92.9 7.1 
65 9.6 23.8 24.3 54.3 44.9 
44 5.4 61.4 22.5 88.0 12.0 
46 4.0 22.8 22.0 74.4 23.5 
57 0.8 55.1 11.7 74.6 24.3 
52 0.7 43.8 24.1 80.0 19.9 
65 2.3 58.9 35.6 90.9 8.9 
57 3.7 60.0 28.9 89.6 9.6 
10 34 2.4 40.4 22.5 73.4 26.4 
11 70 0.9 46.3 24.7 66.2 32.4 
12 49 1.0 49.4 17.8 78.2 20.5 
13 30 2.3 53.5 4.2 94.6 1.6 
14 55 0.7 28.5 27.9 67.6 19.4 
15 45 3.0 9.5 5.4 71.6 26.9 
16 48 0.9 35.5 28.6 73.0 27.0 
Median 48.5 2.1 45.1 23.3 74.5 22.0 

CD19/CD22 dual-targeted CAR T-cell expansion and evaluation of systemic inflammatory markers

The CD19/CD22 dual-targeted CAR T–cell count in the peripheral blood (PB) was detected and assessed by flow cytometry. As shown in Fig. 3A, the CAR T cells in the PB expanded in all patients. The median peak CAR T-cell percentage among total CD3+ T cells was 40.6% (95% CI, 16.8%–54.9%). The median peak of CAR T cells among total CD3+ T cells was 45.5% (95% CI, 12.0%–64.9%) in patients achieving CR versus 34.4% (95% CI, 3.2%–53.3%) in those not achieving CR; this difference did not reach statistical significance, likely because of the low number of patients (P = 0.336; Fig. 3B). We also assessed a panel of cytokines previously reported as being relevant to CAR T-cell therapy using patient serum samples (21). As shown in Fig. 3C, no significant changes in IL2, IL6, IL7, IL15, granzyme B, GM-CSF, MCAP-1, IFNγ, and TNFα were detected in patient serum at initiation (1–2 days after CD19/CD22 dual-targeted CAR T-cell infusion), peak and recovery of CRS following CAR T-cell infusion. Granzyme B levels correlated with the peak CRS stage, but no other cytokines showed clinical correlations.

Figure 3.

CAR T-cell expansion and cytokine/chemokine serum levels. A, Kinetics of individual CAR T-cell expansion assessed as percentage of CAR T cells out of total peripheral blood CD3+ T cells on the indicated days. B, Correlation between CAR T-cell expansion and clinical overall response. CAR T-cell frequency in patients achieving CR and those not achieving CR are depicted. The median peak percentages of CAR T cells in patients achieving CR was 45.5% (95% CI, 12.0%–64.9%) versus 34.4% (95% CI, 3.2%–53.3%) in those not achieving CR (P = 0.336). C, Levels of cytokines and chemokines in serum in correlation with development of CRS (IL2, IL5, IL6, IL7, IL15, granzyme B, GM-CSF, MCP-1, IFNγ, and TNFα) at initiation (1–2 days after CD19/CD22 dual-targeted CAR T-cell infusion), peak, and recovery of CRS following CAR T-cell infusion. ns, not significant. *, **, ***, and **** stand for P < 0.05, P < 0.01, P < 0.001, and P < 0.0001, respectively.

Figure 3.

CAR T-cell expansion and cytokine/chemokine serum levels. A, Kinetics of individual CAR T-cell expansion assessed as percentage of CAR T cells out of total peripheral blood CD3+ T cells on the indicated days. B, Correlation between CAR T-cell expansion and clinical overall response. CAR T-cell frequency in patients achieving CR and those not achieving CR are depicted. The median peak percentages of CAR T cells in patients achieving CR was 45.5% (95% CI, 12.0%–64.9%) versus 34.4% (95% CI, 3.2%–53.3%) in those not achieving CR (P = 0.336). C, Levels of cytokines and chemokines in serum in correlation with development of CRS (IL2, IL5, IL6, IL7, IL15, granzyme B, GM-CSF, MCP-1, IFNγ, and TNFα) at initiation (1–2 days after CD19/CD22 dual-targeted CAR T-cell infusion), peak, and recovery of CRS following CAR T-cell infusion. ns, not significant. *, **, ***, and **** stand for P < 0.05, P < 0.01, P < 0.001, and P < 0.0001, respectively.

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B-cell aplasia

CD19+ B cells in PB were detected by flow cytometry to monitor patients for the development of B-cell aplasia, which can be used as an indirect biomarker of CAR T-cell function. B-cell aplasia occurred in all the patients and persisted for up to 3 to 24 months after CAR T-cell infusion (Supplementary Fig. S2).

OR rates, OS, PFS, and nonrelapse mortality

With a median follow-up of 397 days, the OR rate, as assessed by PET/CT 1 month after CAR T-cell infusion, was 87.5% (14/16); 62.5% (10/16) achieved CR and 25.0% (4/16) achieved PR (Fig. 4A and B). OR rates were the same at 3 months after CAR T-cell infusion. Median OS for the entire group was not reached. OS rates at 1 and 2 years were 77.3% and 77.3%, respectively (Fig. 4C). Median PFS was 246 days. One- and 2-year PFS rates were 40.2% and 40.2%, respectively (Fig. 4D). The 1-year DOR was 45.9% (Fig. 4E), and the median DOR was 210 days. The 2-year DOR in patients who achieved CR was significantly higher than that in patients who did not achieve CR (66.7% and 0.0%, respectively; P = 0.007; Fig. 5A). Notably, the 2-year OS in patients who achieved CR was significantly higher than that in patients who did not achieve CR (100% and 41.7%, respectively; P = 0.015; Fig. 5A). Similar to the OS rates, the 2-year PFS in patients who achieved CR was significantly higher than that in patients who did not achieve CR (66.7% and 0.0%, respectively; P < 0.001; Fig. 5A).

Figure 4.

Clinical outcome of patients following the infusion of CD19/CD22 dual-targeted CAR T cells. A, Treatment response of each patient after CD19/CD22 dual-targeted CAR T-cell treatment and the DOR. Patient number is shown to the left. B, PET/CT imaging pre- and post-CAR T-cell therapy (Patient 10 and Patient 15 in Table 1) are shown. C–E, Kaplan–Meier curves of 1- and 2-year OS, PFS, and DOR.

Figure 4.

Clinical outcome of patients following the infusion of CD19/CD22 dual-targeted CAR T cells. A, Treatment response of each patient after CD19/CD22 dual-targeted CAR T-cell treatment and the DOR. Patient number is shown to the left. B, PET/CT imaging pre- and post-CAR T-cell therapy (Patient 10 and Patient 15 in Table 1) are shown. C–E, Kaplan–Meier curves of 1- and 2-year OS, PFS, and DOR.

Close modal
Figure 5.

Clinical outcome (OS, PFS, and DOR) in correlation with overall response to the dual-targeted CAR T cells and number of previous lines of therapy. A, Kaplan–Meier curves of OS, PFS, and DOR in patients achieving CR versus not achieving CR. B, Kaplan–Meier curves of OS, PFS, and DOR in patients receiving two and more than two prior lines of chemotherapy.

Figure 5.

Clinical outcome (OS, PFS, and DOR) in correlation with overall response to the dual-targeted CAR T cells and number of previous lines of therapy. A, Kaplan–Meier curves of OS, PFS, and DOR in patients achieving CR versus not achieving CR. B, Kaplan–Meier curves of OS, PFS, and DOR in patients receiving two and more than two prior lines of chemotherapy.

Close modal

With regard to previous lines of chemotherapy, PFS and DOR were significantly better in patients who had previously received two lines of chemotherapy compared with those who had received more than 2 lines; 1-year PFS rates were 68.6% and 16.7%, respectively (P = 0.049), and 1-year DOR was achieved in 80.0% and 18.8% of patients, respectively (P = 0.014; Fig. 5B). No differences were observed in OS between patients who had previously received two lines of chemotherapy versus those who had received more than 2 lines of chemotherapy; 1-year OS rates were 83.3% and 71.1%, respectively (P = 0.613; Fig. 5B). The nonrelapse mortality rate was 0%. One patient with stable disease (patient no. 5), one patient with progressive disease (patient no. 11), and one patient with partial remission (patient no. 2) eventually died of disease progression (Fig. 4A). Of the 16 patients recruited, none died of CAR T-cell–associated toxicities. There were no significant differences in response rates between patients in the POC study and the clinical trial (Supplementary Table S3).

Relapse

Three patients (Nos. 6, 12, and 15) relapsed 8, 6, and 4 months after CAR T–cell infusion, respectively. Patient No. 6 suffered from visual disturbance and was considered to have retrobulbar relapse by CT scan. The patient refused a mass biopsy and received irradiation, which led to CR again, and the patient was still alive at data collection. Patient no. 12 had localized intraparenchymal lesions and underwent lumbar puncturel CD19+/CD22+ lymphoma cells were detected in the CSF by flow cytometric analysis. The patient then received ibrutinib, lenalidomide, and dexamethose treatment, which led to PR, and the patient was still alive at data collection. Patient no. 15 had localized intraparenchymal lesions and underwent biopsy; CD19+/CD22+ lymphoma cells were detected by flow cytometry. The patient received irradiation, which led to CR again.

Multivariate analyses of predictive factors for OS, PFS, and DOR

We performed multivariate analysis of predictive factors for outcomes. Prior lines of chemotherapy (n > 2; HR = 135.784; 95% CI, 1.069–17248.110; P = 0.047) and best OR (non-CR; HR = 0.046; 95% CI, 0.006–0.358; P = 0.003) were two independent prognostic factors associated with poor PFS (Table 2; Supplementary Table S4). The best OR (non-CR; HR = 0.087; 95% CI, 0.013–0.600; P = 0.013) and prior lines of chemotherapy (HR = 4.807; 95% CI, 1.373–16.830; P = 0.014) were also independent prognostic factors associated with DOR (Table 2; Supplementary Table S5). No factors were predictive for OS (Table 2; Supplementary Table S6). Baseline lactate dehydrogenase, disease stage at diagnosis, International Prognosis Index (IPI), bone marrow involvement, patient age, CAR T-cell dose, and CRS grade were not significant prognostic factors for any outcome parameter (Supplementary Tables S4–S6).

Table 2.

Multivariable analyses for factors impacting PFS, DOR, and OS in patients who received CD19/CD22 dual-targeted CAR T-cell therapy.

FactorsHR (95% CI)P value
PFS 
 Prior lines of chemotherapy  135.784 (1.069–17248.110) 0.047 
 Best OR CR vs. non-CR 0.017 (0.000–0.935) 0.046 
 IL1a at baseline  2.066 (0.992–4.304) 0.053 
DOR 
 Best OR CR vs. non-CR 0.087 (0.013–0.600) 0.013 
 Prior lines of chemotherapy  4.807 (1.373–16.830) 0.014 
OS 
 IL15 at baseline  1.052 (0.991–1.117) 0.095 
FactorsHR (95% CI)P value
PFS 
 Prior lines of chemotherapy  135.784 (1.069–17248.110) 0.047 
 Best OR CR vs. non-CR 0.017 (0.000–0.935) 0.046 
 IL1a at baseline  2.066 (0.992–4.304) 0.053 
DOR 
 Best OR CR vs. non-CR 0.087 (0.013–0.600) 0.013 
 Prior lines of chemotherapy  4.807 (1.373–16.830) 0.014 
OS 
 IL15 at baseline  1.052 (0.991–1.117) 0.095 

Note: P values < 0.05 were considered statistically significant and appear in bold.

Adverse events and toxicities of CD19/CD22 dual-targeted CAR T-cell therapy

All patients developed CRS, albeit mostly low-grade (grades 1–2), and responded to conventional therapy (Supplementary Table S7). Eleven had grade 2 CRS, which manifest as fever, mild tachypnea, and mild hypotension. Four patients experienced grade 1 CRS, and only one patient had grade 4 CRS (high fever, dyspnea, and severe hypotension); this patient responded to established conventional therapies and was the only study patient who received tocilizumab (anti-IL6). We observed no grade 5 CRS, and no patients died due to CRS or CAR T-cell–related toxicities. The median day of CRS onset was 2.5 days (range, 1–10 days) after CAR T-cell infusion, and CRS lasted for a median of 5.9 days (range, 2–9 days).

No patients developed CAR T-cell–related neurologic toxicities. The most frequent treatment-emergent adverse events were fever, fatigue, headache, and diarrhea, observed in 100% of patients (Supplementary Table S7). Other adverse events included hypotension (43.8%), hypoxia (43.8%), cough (75.5%), nausea (93.8%), chills (87.5%), vomiting (81.3%), and weight loss (87.5%); these were fully reversible with supportive care. Laboratory findings included hypoalbuminemia (87.5%), anemia (81.3%), decreased platelet count (68.8%), febrile neutropenia (68.8%), decreased lymphocyte count (56.3%), decreased neutrophil count (68.8%), and pleural effusion (43.8%; Supplementary Table S7).

Various studies have shown that CD19-targeted CAR T cells are an attractive and effective therapy for R/R B-cell lymphoma (3). However, the relatively low CR rate and CD19-negative relapse have emerged as major challenges for long-term disease-free survival after CD19-targeted CAR T-cell therapy. Here, we have reported a clinical trial of CD19/CD22 dual-targeted CAR T cells as a treatment for patients with R/R aggressive B-cell lymphoma. The generation and manufacturing of the CD19/CD22 dual-targeted CAR T cells was uneventful and successful in all study patients. Moreover, the dual-targeted CAR T cells resulted in encouraging clinical responses and sustained CRs in patients with R/R aggressive B-cell lymphoma; none of the patients showed CD19-negative relapse. Notably, the incidence and severity of CAR T-cell–associated CRS were low, and no patients developed neurotoxicities. Other toxicities and side effects were infrequent.

The successful generation and manufacturing of CAR T cells are the basis of CAR T-cell therapy. Many factors affect CAR T-cell engineering. The size of the CAR construct may affect the level of CAR expression on the T-cell surface. Large constructs and long gene sequences are usually accompanied by low genetic transduction efficiency (22). In this clinical trial, the transduction rate of the CD19/CD22 dual-targeted CAR T cells was high. This transduction yield is consistent with the yield reported for monospecific CD19-targeted CAR T cells (4). The molecular characteristics of a CAR construct, including scFv source, scFv order, linker length, spatial considerations, and CAR configuration, also have a major impact on the potency of CAR T cells. Our CD19/CD22 dual-targeted CAR T–cell construct seemed to be highly efficient. In comparison with commonly used monospecific CD19-targeted CAR T cells, the cytotoxicity of the CD19/CD20 dual-targeted CAR T cells in K562 cells at an E:T ratio of 1:1 is about 20% (23). Moreover, the cytokine profile of the CD19/CD22 dual-targeted CAR T cells was similar to that of monospecific CD19-targeted CAR T cells, and the dual-targeted cells showed superior target-cell lysis when challenged with optimal target cells.

In previous studies, approximately 50% of treated patients with diffuse large B-cell lymphoma obtained CRs with monospecific CD19-targeted CAR T cells (24, 25). Most studies of such cells evaluate treatment responses and whether CR is achieved 3 months following CAR T-cell infusion (26). Notably, achievement of CR or PR 3 months after CAR T-cell infusion is predictive of the long-term durability of clinical response (4). In the current study, response at 3 months after CAR T-cell infusion did not differ from the response at 1 month. At 1 month after CAR T-cell infusion, 14 of 16 (87.5%) patients achieved clinical response, and 10 of 16 (62.5%) achieved CR, indicating rapid antilymphoma effects of the CD19/CD22 dual-targeted CAR T cells. The CD19/CD22 dual-targeted CAR T cells also induced durable responses that lasted more than 2 years, with no need for consolidation therapy, including stem cell transplantation, with the similar results to those reported for the monospecific CD19-targeted CAR T cells in the ZUMA1 and Juliet trials (27, 28).

Sequential infusion of monospecific CD19-targeted and monospecific CD22-targeted CAR T cells in 38 patients with B-cell lymphoma led to OR and CR rates of 72.2% and 50.0%, respectively, in one study (29). In another study, sequential monospecific CD22-targeted CAR T cells and monospecific CD22-targeted CAR T cells to patients with R/R aggressive B-cell lymphoma of the gastrointestinal tract yielded an OR rate of 76.9% and a CR rate of 53.8% (30). In addition, administration of CD19/CD22 dual-targeted CAR T cells to patients with R/R ALL shows potent antileukemic activity and a good safety profile (31, 32). The results achieved with our CD19/CD22 dual-targeted CAR T cells in patients with advanced lymphoma were similar to those observed by sequential administration of monospecific CD19-targeted and monospecific CD22-targeted CAR T cells and those reported with CD19/CD22 dual-targeted CAR T cells in patients with ALL.

For monospecific CD19 CAR T-cell therapy, CD19-negative relapse has emerged as the most significant obstacle for long-term disease-free survival (8, 9, 33, 34). In our study of CD19/CD22 dual-targeted CAR T cells, once most patients had achieved CR, the CR was sustained for up to 21.6 months. Only three patients relapsed, and no patients showed antigen-loss relapse, which may support the concept that dual antigen–targeted CAR T cells may overcome the limitation of antigen-loss escape observed for monospecific CAR T cells because the risk of dual-antigen loss is low.

Although no factors were found to be associated with OS, the number of prior lines of chemotherapy and best OR were two independent risk factors associated with both PFS and DOR. Administration of CD19/CD22 dual-targeted CAR T cells to patients receiving two of fewer lines of previous chemotherapy may be the most effective approach and should be explored in greater detail in the future.

Importantly, the CD19/CD22 CAR T-cell treatment regimen was generally well tolerated with very few side effects; mild and transient mostly grade 1/2 CRS was observed, with only one patient developing grade 4 CRS, and no neurotoxicities were found. This safety profile is remarkable in comparison with previously reported CAR T-cell studies. For example, using ASTCT CRS grading scale, one study reported that 13.5% of patients with refractory large B-cell lymphoma experienced grade 3 CRS or higher receiving tisagenlecleucel treatment (35) and another reported 7 (14.5%) patients with R/R diffused large B-cell lymphoma had grade 3 or higher CRS after monospecific CD19-targeted CAR T-cell treatment (36). Similar with our study, Shah and colleagues reported that CD19/CD20 dual targeted CAR-T cell therapy for R/R diffuse large B-cell lymphoma, grade 3 CRS or higher was observed in 5% of patients (37). The underlying mechanisms need further clarification. In addition, the level of cytopenia we observed was similar to that reported with monospecific CD19-targeted CAR T cells, despite the increased potency of the CD19/CD22 dual-targeted CAR. Less than 20% of patients in our study had grade 3 or worse cytopenia, no platelet or red blood cell transfusion was required, and blood counts usually normalized by 1 month after CAR T-cell infusion.

The low rate of toxicity of the CD19/CD22 dual-targeted CAR may be related to the low levels of inflammatory cytokines induced, as assessed using patient serum samples after CAR T-cell infusion; the levels of inflammatory cytokines and immunomodulators were similar to the observed basal level, with only mild increases. Thus, the high potency and cytotoxicity we observed both in vitro and in vivo with the dual CD19/CD22 dual-targeted CAR T cells may be mediated by high granzyme B secretion, as detected in patient serum after infusion.

Another practical aspect of this approach is a reduction in manufacturing cost. The production cost of the dual CD19/CD22 dual-targeted CAR T cells was lower than that of two monospecific CAR T cells for sequential infusion CAR T-cell therapy. For the dual CD19/CD22 dual-targeted CAR T cells, both CD19 and CD22 scFvs were engineered on the same CAR construct; thus, the production cost was reduced. Moreover, fewer T cells were required for dual-antigen CAR T-cell generation, providing a practical clinical advantage, particularly in heavily treated patients with R/R aggressive B-cell lymphoma with severe leukopenia or neutropenia; indeed, this approach may enable sufficient T-cell collection via leukapheresis, which can be a problem in these patients.

There were several limitations to the current study, including the lack of randomized, case-controlled, multicenter, prospective data and the limited sample size. These limitations may have affected the reliability of the statistical analysis. However, these limitations are typical of early-stage, first-in-human trials, and our study has demonstrated the feasibility and safety of dual CD19/CD22 dual-targeted CAR T-cell therapy for patients with advanced R/R aggressive B-cell lymphoma. The dual CD19/CD22 dual-targeted CAR T cells proved to be potent and exhibit impressive antilymphoma activity with minimal toxicity and only mild-to-moderate CRS. Based on our study results, it is conceivable that dual-antigen or multitargeted CAR T-cell therapy may overcome the shortcomings of monospecific CD19-targeted T-cell therapy, improving the chance of curing R/R B-cell malignancies. Further studies in well-designed multicenter trials, including a formal comparison of CD19/CD22 dual-targeted CAR T cells with monospecific CD19- or CD22-targeted CAR T cells and/or sequential monospecific CD19- and CD22-targeted CAR T-cell administration, are required in order to establish CD19/CD22 dual-targeted CAR T-cell immunotherapy as a backbone for future therapy in patients with R/R aggressive B-cell lymphoma.

A.H. Chang is a founding member of Shanghai YaKe Biotechnology Ltd., a biotechnology company focusing on research and development of tumor cellular immunotherapy. No disclosures were reported by the other authors.

G. Wei: Conceptualization, data curation, formal analysis, validation, investigation, methodology, writing–original draft, writing–review and editing. Y. Zhang: Validation, investigation, methodology, writing–original draft. H. Zhao: Conceptualization, investigation, methodology, writing–original draft. Y. Wang: Formal analysis, investigation, methodology, writing–original draft. Y. Liu: Data curation, writing–review and editing. B. Liang: Investigation, methodology, writing–original draft, writing–review and editing. X. Wang: Investigation, methodology, writing–original draft, writing–review and editing. H. Xu: Investigation, visualization, methodology, writing–original draft, writing–review and editing. J. Cui: Formal analysis, investigation, methodology, writing–original draft. W. Wu: Investigation, methodology, writing–original draft, writing–review and editing. K. Zhao: Validation, investigation, methodology, writing–original draft. A. Nagler: Methodology, writing–original draft, writing–review and editing. A.H. Chang: Investigation, methodology, writing–original draft, project administration, writing–review and editing. Y. Hu: Conceptualization, data curation, software, formal analysis, supervision, funding acquisition, investigation, writing–original draft. H. Huang: Supervision, funding acquisition, validation, investigation, visualization, methodology, writing–original draft, project administration, writing–review and editing.

This study was supported by the National Natural Science Foundation of China (grant numbers 81730008, 81770201, and 81870153) and the Key Project of Science and Technology Department of Zhejiang Province (grant numbers 2019C03016, 2018C03016-2, and 2021C03010).

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.

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Supplementary data