Chimeric antigen receptor T cells persist and induce sustained remissions in relapsed refractory chronic lymphocytic leukemia

CTL019 cell manufacturing feasibility and product characteristics

Product characteristics are outlined in table S1. All but one infused patient met the target manufactured dose range of 1.5 × 10⁷ to 5 × 10⁹ total nucleated cells (TNCs). Patients received a median of 7.5 × 10⁸ (range, 1.7 × 10⁸ to 50 × 10⁸) TNCs. CTL019 cells refer to the transduced, CAR-expressing T cells; the median transduction efficiency of the 14 manufactured products was 20.1% (range, 4.7 to 39.2%) and the total CTL019 dose infused was a median of 1.6 × 10⁸ cells (range, 0.14 × 10⁸ to 11 × 10⁸) infused over days 0, 1, and 2. Eight (57%) patients completed the full 3-day split-dose regimen; three patients received only one fraction (10% of the manufactured dose) and three patients received only two fractions (total of 40% of the manufactured dose) after developing fevers within 24 hours of CTL019 infusion, accounting in part for the variability in cell dose. The total infused product contained a median of 81% (range, 68 to 95%) of CD4-positive cells and a median of 20% (range, 4.6 to 34%) of CD8-positive cells.

Products from 12 patients were tested for reactivity against CD19-positive targets using functional flow cytometry, measuring degranulation as a measure of cytolytic activity, and the intracellular accumulation of cytokines. All 12 products showed cytolytic activity against the CD19 targets and produced high levels of most cytokines, indicating that the CTL019 cells generated for this patient cohort expressed a functional CAR (fig. S2).

Patient outcomes: Response and survival

Response was determined based on the International Workshop Group on CLL response criteria (13). In addition, deep sequencing analysis of the immunoglobulin heavy chain (IGH) locus was performed for each patient and used for disease detection in blood and bone marrow over time. The best overall response and clinical outcomes are summarized in Table 2. The median follow-up for all 14 treated patients is 19 months (range, 6 to 53 months). Eight of 14 [57%; exact 95% confidence interval (CI), 29 to 82%) patients responded. Four patients (29%) achieved CR. No patient with CR has relapsed with median duration of response of 40 months (range, 21 to 53 months from time of infusion). Remissions included eradication of minimal residual disease (MRD) as assessed through deep sequencing analysis which failed to identify the presence of CLL clones in the blood or bone marrow from subjects 01, 02, and 09 by month 1 and from subject 10 by month 3 [Table 3, table S2, and (8)]. Three of these patients remain alive with no evidence of recurrent disease 28 to 53 months after CTL019 infusion. Subject 09 was in CR and underwent removal of basal cell carcinoma from his lower leg 15 months after CTL019 infusion; the surgical site became infected leading to lethal pseudomonas sepsis and ecthyma gangrenosum 21 months after CTL019 infusion.

Four subjects (03, 05, 12, and 22) (29%) achieved a partial response (PR) within the first month of CTL019 infusion. The median duration of response in the four patients achieving PR was 7 months (range, 5 to 13 months). In two of these patients (03 and 05), there was improvement in lymphocytosis, marrow involvement, and adenopathy, but both patients progressed 4 months after the infusion. Subject 12 entered treatment with extensive marrow infiltration, transfusion dependence, and diffuse bulky adenopathy and achieved PR. After infusion, he had a 2-log CTL019 expansion with complete clearance of CLL from his blood and marrow and >50% reduction in his adenopathy. The patient died of complications from a massive pulmonary embolus 6 months after T cell infusion with no evidence of CLL in the blood or marrow. At autopsy however, sampling of residual enlarged abdominal lymph nodes showed residual infiltration with CD19⁺ CLL. Subject 22 also had extensive marrow infiltration, circulating disease, and bulky adenopathy at baseline. After CTL019 infusion, he achieved PR with complete clearance of CLL from the blood and marrow with >50% reduction in adenopathy by 3 months and sustained at 6 months. Nine months after infusion, he presented with rapidly growing adenopathy; lymph node biopsy showed diffuse large B cell lymphoma arising from CLL (Richter transformation) with cells that were CD19-dim. Marrow biopsy showed about 15% involvement with CLL and large transformed-appearing lymphocytes. At the time of transformation and relapse, no CTL019 cells were detected either in his blood, marrow, or lymph node biopsy specimen by flow cytometry though CTL019 cells were detected by quantitative polymerase chain reaction (qPCR) in the blood and bone marrow at the time of progression.

Six subjects (43%) had no response and all six progressed within 1 to 9 months (median, 4 months) of CTL019 therapy. T cell expansion was limited in all six of these subjects (median, 420 copies/μg; range, 6.5 to 13,876, or 0.2% of the total CD3⁺ T cells). CLL cells from these patients were not resistant to lysis in vitro. Two of these subjects have died; one from complications of CLL and infection 8 months after CTL019, and one from complications of allogeneic stem cell transplantation performed 9 months after CTL019 and after two additional therapies (ofatumumab and CHOP) for his CLL. The other four subjects are alive 9 to 15 months after CTL019, all receiving alternative therapies.

Leukemia response to CTL019 was assessed using IGH deep sequencing in all 14 patients (Table 3 and table S2) and four patterns were identified: (i) CTL019 infusion was followed by eradication of MRD with rapid and persistent disappearance of the leukemic clone (subjects 01, 02, and 09). Peripheral blood DNA from subjects 01, 02, and 09 was sequenced up to 4, 2, and 1 year, respectively, after infusion, and the leukemic clone was undetectable in all cases; (ii) CTL019 infusion resulted in a delayed complete eradication of MRD with molecular remission determined by deep sequencing 3 months after infusion; (iii) Some patients responded with a transient complete (subject 22) or incomplete (subjects 03, 05, 12, 17, and 25) disappearance of the leukemic clone; (iv) CTL019 did not result in any decrease of the number of IGH reads or leukemic clone in subjects 06, 07, 14, and 18.

The estimated median overall survival for the 14 treated and evaluable subjects was 29 months, and the 18-month overall survival was 71% (95% CI, 40.6 to 88.2%). The estimated median progression-free survival (PFS) for treated and evaluable patients was 7 months, and the 18-month PFS was 28.6% (95% CI, 8.8 to 52.4%) (Fig. 1 and table S6). Whereas one subject in CR died 21 months after infusion because of complications related to infection, no patient who achieved CR progressed. All other patients ultimately progressed or died of disease-related complications 1 to 10 months after infusion.

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Fig. 1

Overall survival and PFS

(A and B) Kaplan-Meier curves for overall survival (A) and PFS (B) were estimated for the 14 subjects who received therapy. Shaded region indicates 95% CI. Number of subjects at risk is presented in a 3-month interval. Vertical tick marks indicate time points with censored data. (Source data, table S6).

As shown in fig. S1, 23 patients were screened for this study and 9 were not treated. Five patients did not proceed from screening to enrollment because they either declined participation or failed the screening. Therefore, a total of 18 patients were enrolled in the study and 4 were not treated because of rapid disease progression and/or death after enrollment, before therapy (n = 3), or they decided to withdraw consent in favor of alternative therapies (n = 1). Because a delay between enrollment and start of therapy is unique to these types of cell therapy approaches, we performed an analysis of PFS of all 18 enrolled subjects. The median PFS from the date that eligibility was confirmed for all 18 subjects was 9 months (95% CI, 3 to 13.6).

There was no statistically significant association between response and patient age, number of previous therapies, stage at enrollment, deletion of chromosome 17p, IGH variable (IGHV) mutation status, or CTL019 cell dose (all P >0.05, table S3). Because all patients had extensive disease, it was not possible to determine an association between disease burden and response. We also analyzed CD19 expression levels on CLL cells before infusion, and there was no difference in CD19 mean fluorescence intensity between responders and nonresponders, suggesting that target density did not predict response.

CTL019 in vivo expansion and persistence

Persistence and quantitation of CTL019 cells (transduced and CAR-expressing) were performed by flow cytometry to identify T cells expressing the anti-CD19 CAR and by qPCR to identify gene-modified T cells. The degree of expansion and the duration of persistence were correlated to response (Fig. 2 and table S7). The expansion of CTL019 cells was coincident with the development of cytokine release syndrome (CRS) in most cases (described below), and CAR T cells expanded to high levels in all responding patients.

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Fig. 2

CTL019 expansion by qPCR in the first 12 months

Peripheral blood CTL019 expansion was measured as copies per microgram of genomic DNA by qPCR in 14 subjects. Values below the quantitative limit of detection (<25 copies/μg DNA) are shown with open circles. (Source data, table S7).

Among the four patients achieving CR, CTL019 peak expansion was 25,070 to 409,645 (median, 73,237) copies/μg by qPCR; for the subjects that had expansion measured by flow cytometry, the peak percentage of CD3⁺ cells that were positive for CTL019 was 34.3 and 81.9% of CD3⁺ in peripheral blood mononuclear cells (PBMCs). Expansion was less robust in four patients achieving PR (range, 1607 to 130,258; median, 33,453 copies/μg) by qPCR or a median of 18.3% of the CD3⁺ population. Expansion was minimal in patients who did not achieve a response (range, 6.5 to 13,876; median, 420 copies/μg) by qPCR or a median of 0.2% of the total CD3⁺ population by flow cytometry. The peak value of CTL019 was statistically associated with response (Fig. 3 and table S8) (P = 0.013 by qPCR; P = 0.008 by flow cytometry).

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Fig. 3

Association between peak CTL019 expansion and response

(A and B) Peak CTL019 expansion in the first 3 months measured by flow cytometry (A) in 11 subjects and qPCR (B) in 14 subjects. Flow cytometry detection was not available for subjects 01, 02, and 03 because an antibody to detect CTL019 cells was not available at the time of their treatment. Wilcoxon rank-sum P values are provided. (Source data, table S8).

Long-term persistence of CTL019 cells has been detected by means of flow cytometry or qPCR to be 14 to 49 months in the four patients who achieved CRs (Fig. 4A and table S9). All four patients had ongoing B cell aplasia at the time last tested for persisting CTL019 cells, suggesting that the persisting cells remain functional by inhibiting the development of CD19⁺ B cells. Furthermore, to directly show that persisting CTL019 cells were functional, CAR-modified T cells were isolated from subject 02 almost 3 years after infusion; the CTL019 cells retained immediate and specific reactivity against CD19⁺ targets, highlighting that at least some of these cells remain functional without exhaustion for years after infusion (Fig. 4, B and C and table S10).

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Fig. 4

Long-term persistence of CTL019 cells and poly-functionality in patients achieving CR

(A) Long-term persistence of CTL019 cells by flow cytometry (solid circle) and qPCR (solid triangle) beyond 12 months from infusion. All observed values were above the limit of detection by flow cytometry (0.1%) and above the limit of quantification by qPCR (<25 copies/μg DNA). (B and C) Purified PBMCs from patient UPCC04409-02 from the indicated postinfusion time points were stimulated for 6 hours with APC-expressing CD19 or control antigen (mesothelin) and examined for de-granulation (CD107a) and cytokine protein expression. (B) Contour plots displaying CD107a versus MIP-1β (top two rows) or IL-2 versus IFN-γ expression (bottom rows) in CD8⁺ CAR⁺ T cells stimulated with K562 cells expressing control antigen (mesothelin) or CD19. (C) Summary graph of functional response of CAR19⁺ CD8 T cells upon stimulation with CD19⁺ target cells over time. Data are plotted from (B) and are depicted as the background-subtracted frequency of this subset. (Source data, tables S9 and S10).

Toxicity

CTL019 infusions were well tolerated. Infusional toxicities were infrequent and mild (grade <2) and included primarily low-grade fevers and chills. No patient had a significant reaction during CTL019 infusion. The most frequent related events were associated with complications of neutropenia (including fevers) and delayed CRS, which correlated with in vivo CTL019 expansion. Two cases of tumor lysis syndrome were noted. One patient died in remission 21 months after T cell infusion, after developing overwhelming ecthyma gangrenosum from a pseudomonas wound infection from a skin biopsy site.

Cytokine release and macrophage activation syndrome

Symptoms from CRS ranged from mild to severe and were frequently self-limiting. CRS presenting symptoms included fever, myalgia, and nausea, and in severe cases, escalated to hypotension, capillary leak, and hypoxia. CTCAE (Common Terminology Criteria for Adverse Events) version 3.0 captures CRS as an acute infusional toxicity but is inadequate to describe delayed CRS that may occur days after the administration of CTL019. Therefore, we devised a novel grading scale for CRS (table S4, A and B). In addition, during CRS, we also noted concurrent cytopenia, coagulopathy, histologic macrophage activation highlighted by CD163 staining, and hemophagocytosis, as noted in one patient with a marrow biopsy at peak of CRS, and patients with CRS had marked elevations in ferritin, C-reactive protein (CRP), and soluble interleukin-2 receptor (sIL-2R), findings consistent with macrophage activation syndrome (MAS) or hemophagocytic lymphohistiocytosis (HLH).

Nine patients developed CRS 1 to 14 days (median, 7 days) after CTL019 infusion, including one grade 1, two grade 2, two grade 3, and four grade 4. CRS required intervention with anti-cytokine directed therapy in five patients a median of 9.5 days after infusion (range, 9 to 55 days), and four patients required an intensive care unit (ICU) level of care for complications related to CRS, such as hypotension and hypoxia, with a median length of ICU stay of 6 days (range, 4 to 9 days).

We previously noted marked elevations in IL-6 associated with severe CRS after CTL019 infusions in a patient with ALL and rapid reversal with the IL-6 receptor–blocking antibody tocilizumab (14). Before this observation, one patient (subject 03) received corticosteroids to successfully reverse CRS. For subsequent patients, tocilizumab was incorporated into the management of severe CRS for CLL. Four patients received tocilizumab (with the addition of steroids in two patients), which resulted in rapid defervescence, stabilization of blood pressure, and rapid improvement in biochemical abnormalities associated with macrophage activation (ferritin and CRP). An example of the biochemical response of ferritin to tocilizumab is illustrated in fig. S3.

There was higher peak expansion of CTL019 associated with CRS, and CRS was associated with clinical response (P < 0.05) as shown in fig. S4. Compared to patients with no or mild CRS (grade 0 to 1), patients with significant CRS (grade 2 to 4) had higher peak levels of IL-6 (median peak, 269 versus 34 pg/ml; P = 0.014). Levels of sIL-2R were higher in patients with grade 2 to 4 CRS (median peak, 10,962 versus 4177 pg/ml; P =0.008), whereas no statistically significant differences were found in serum interferon-γ (IFN-γ) or IL-2 (Fig. 5 and table S11). The data were limited in the month after the infusion for ferritin (eight subjects) and CRP (five subjects), which prevented a definitive analysis of their association with CRS grade. Patients with grade 2 to 4 CRS had higher peak levels of ferritin (median, 21,502 versus 1980) and C-reactive protein (median, 160 versus 42) than patients with grade 0 to 1 CRS; because of the small number of patient samples available, detailed statistical analysis was not performed, but this limited data set suggests that ferritin and/or CRP may serve as useful biomarkers for CRS activity and response to treatment. There was no disease or patient-specific characteristics that predicted the development of CRS including age, previous therapy, genetic risk profile, or cell dose (table S5).

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Fig. 5

Peak cytokines values over the first month after infusion

(A to D) Peak values for IL-6 (A), IFN-γ (B), IL-2 receptor antagonist (IL-2RA) (C), and IL-2 (D) by CRS grades are displayed. Gray symbols represent nonresponders; black symbols are those who achieved a complete or partial remission. Individual values are plotted with a symbol that reflects their CRS grade (1 to 4). Wilcoxon rank-sum P values are also provided. Cytokine values were missing for one grade 4 subject who achieved CR (n = 13). (Source data, table S11).

All four recipients of tocilizumab for CRS had ongoing T cell expansion at the time of tocilizumab infusion. After tocilizumab, there was marked improvement in symptoms, although in at least two cases, T cell proliferation peaked several days after administration, suggesting that tocilizumab may not fully inhibit ongoing proliferation. Because all patients had persisting cells, it is clear that tocilizumab did not affect long-term T cell survival.

Concurrent with CRS were six recorded neurologic events in five patients that included grade 1 or 2 hallucinations, confusion, or delirium typically associated with high fevers, ICU care, or medication use. There was one case of grade 4 confusion lasting 2 days and attributed at least in part to T cell therapy.

Vector production

The CD19-BB-z transgene (GeMCRIS 0607-793) was designed and constructed as described (27). A lentiviral vector was produced according to current manufacturing practices with a three-plasmid production approach at either Lentigen Corporation (28) or Children’s Hospital of Philadelphia.

T cell collection and generation of CTL019 cells

Autologous T cells were collected by leukapheresis and were stimulated with paramagnetic polystyrene beads coated with anti-CD3 and anti-CD28 monoclonal antibodies (mAbs) (29, 30). Cells were transduced with a lentiviral vector encoding anti-CD19 scFv linked to 4-1BB and CD3-z signaling domains as described (31) and were expanded ex vivo for 10 to 12 days. The median transfection efficiency was 20.1% (range, 4.7 to 39.2%). Release criteria have been previously described (8).

Response assessment

Disease response assessments were done at protocol-defined time points and were based on the International Workshop Group on CLL response criteria (13) with the incorporation of independent radiology review. In addition, investigational assessments for MRD using deep sequencing analysis were performed (see below).

Correlative studies

Sample processing, flow cytometry, cytokine and cytokine receptor analysis, and qPCR analyses were performed as described (11). In addition, the following studies were performed.

Polychromatic flow cytometry

CTL019 administration

Lymphodepleting chemotherapy intended for depletion of T lymphocytes was timed to end about 4 days before infusion of CTL019. Cells were administered by intravenous infusion with a 3-day split-dose regimen (10% on day 0, 30% on day 1, and 60% on day 2) with a total targeted dose of 1 × 10⁹ to 5 × 10⁹ CD3⁺ cells and a minimum acceptable dose of 1.5 × 10⁷ CTL019 cells. Day 1 or 2 infusions were held for fevers or other toxicity. If less than the target dose was available, patients received the total number of manufactured cells as a 3-day split infusion. According to the original protocol design, patients could receive an additional dose of T cells if available on day 10; only one patient (subject 03) received this dose. Subsequently, no additional planned day 10 infusions were given. Subjects were assessed for toxicity planned intervals (a minimum of weekly for 1 month, monthly for 6 months, and then every 3 months) with response assessments performed at 1 month, 3 months, and then every 3 months.

Toxicity grading

Toxicities after CTL019 infusion were graded according to Common Toxicity Criteria (CTC) version 3.0. CTC grading for CRS was designed to grade acute infusional reactions typically seen with antibody therapies and was inadequate to describe the delayed and ongoing CRS from cellular therapy (21); we developed a modified grading scale for CRS designed to describe the severity of the reaction and help identify the need for medical intervention. The revised grading scale for CRS is shown in table S4, A and B.

We thank L. Lledo, H. McConville, J. Gilmore, J. Capobianchi, C. Jemison, M. Harvey, H. Difilippo, E. Veloso, and A. Marshall for clinical research assistance; A. Bersenev, A. Lamontagne, A. Malykhin, N. Manvar, M. O’Rourke, M. Suhoski-Davis, M Kalos, and the members of the Clinical Cell and Vaccine Production Facility for cell manufacturing and testing; Y. Mahnke and L. Tian for correlative sciences contributions; J. Finklestein, I. Kulikovskaya, F. Nazimuddin, M. Gupta, E. Pequignot, J.-M. Navenot, F. Chen, H. Parakandi, M. Bogush, Y. Tanner, and N. Kengle for research support; P. Wood, M. Litchman, A. Quintas, and G. Shah from Novartis and S. Maude from the Children’s Hospital of Philadelphia for discussions and protocol support; B. Jena and L. Cooper (MD Anderson Cancer Center) for provision of the CAR anti-idiotype detection reagent; and B. Fuchs and C. Candeloro for assistance in developing our CRS grading scale and defining pressor doses. We would also like to acknowledge the Data and Safety Monitoring Board, the outstanding clinical support provided by our hematology-oncology faculty and nurses, the other critical care faculty and nurses, and our residents and fellows at the Hospital of the University of Pennsylvania. Most importantly, we acknowledge the courageous participation of all the patients in this trial.

Funding: This study was supported in part by a grant from Novartis and by grants from the Leukemia and Lymphoma Society (Specialized Center of Research Award) and NIH (1R01CA165206 to D.L.P. and C.H.J.; 1K24 CA117879 to D.L.P.); R01CA102646 and R01CA116660 to S.A.G.).