Biology of Blood and Marrow Transplantation
Volume 16, Issue 9 , Pages 1245-1256, September 2010

Antitransgene Rejection Responses Contribute to Attenuated Persistence of Adoptively Transferred CD20/CD19-Specific Chimeric Antigen Receptor Redirected T Cells in Humans

  • Michael C. Jensen

      Affiliations

    • Department of Cancer Immunotherapeutics and Tumor Immunology, Beckman Research Institute, City of Hope National Medical Center, Duarte, California
    • Department of Hematology and Hematopoietic Cell Transplant, Beckman Research Institute, City of Hope National Medical Center, Duarte, California
    • Corresponding Author InformationCorrespondence and reprint requests: Michael C. Jensen, MD, Department of Cancer Immunotherapeutics and Tumor Immunology, Beckman Research Institute, City of Hope National Medical Center, 1500 East Duarte Road, Duarte, CA 91010-3000.
    • ,
  • Leslie Popplewell

      Affiliations

    • Department of Hematology and Hematopoietic Cell Transplant, Beckman Research Institute, City of Hope National Medical Center, Duarte, California
    • ,
  • Laurence J. Cooper

      Affiliations

    • Department of Hematology and Hematopoietic Cell Transplant, Beckman Research Institute, City of Hope National Medical Center, Duarte, California
    • ,
  • David DiGiusto

      Affiliations

    • Department of Hematology and Hematopoietic Cell Transplant, Beckman Research Institute, City of Hope National Medical Center, Duarte, California
    • ,
  • Michael Kalos

      Affiliations

    • Department of Cancer Immunotherapeutics and Tumor Immunology, Beckman Research Institute, City of Hope National Medical Center, Duarte, California
    • ,
  • Julie R. Ostberg

      Affiliations

    • Department of Cancer Immunotherapeutics and Tumor Immunology, Beckman Research Institute, City of Hope National Medical Center, Duarte, California
    • ,
  • Stephen J. Forman

      Affiliations

    • Department of Cancer Immunotherapeutics and Tumor Immunology, Beckman Research Institute, City of Hope National Medical Center, Duarte, California
    • Department of Hematology and Hematopoietic Cell Transplant, Beckman Research Institute, City of Hope National Medical Center, Duarte, California

Received 26 January 2010; accepted 11 March 2010. published online 22 March 2010.

Article Outline

Immunotherapeutic ablation of lymphoma is a conceptually attractive treatment strategy that is the subject of intense translational research. Cytotoxic T lymphocytes (CTLs) that are genetically modified to express CD19- or CD20-specific, single-chain antibody–derived chimeric antigen receptors (CARs) display HLA-independent antigen-specific recognition/killing of lymphoma targets. Here, we describe our initial experience in applying CAR-redirected autologous CTL adoptive therapy to patients with recurrent lymphoma. Using plasmid vector electrotransfer/drug selection systems, cloned and polyclonal CAR+ CTLs were generated from autologous peripheral blood mononuclear cells and expanded in vitro to cell numbers sufficient for clinical use. In 2 FDA-authorized trials, patients with recurrent diffuse large cell lymphoma were treated with cloned CD8+ CTLs expressing a CD20-specific CAR (along with NeoR) after autologous hematopoietic stem cell transplantation, and patients with refractory follicular lymphoma were treated with polyclonal T cell preparations expressing a CD19-specific CAR (along with HyTK, a fusion of hygromycin resistance and HSV-1 thymidine kinase suicide genes) and low-dose s.c. recombinant human interleukin-2. A total of 15 infusions were administered (5 at 108cells/m2, 7 at 109cells/m2, and 3 at 2 × 109cells/m2) to 4 patients. Overt toxicities attributable to CTL administration were not observed; however, detection of transferred CTLs in the circulation, as measured by quantitative polymerase chain reaction, was short (24 hours to 7 days), and cellular antitransgene immune rejection responses were noted in 2 patients. These studies reveal the primary barrier to therapeutic efficacy is limited persistence, and provide the rationale to prospectively define T cell populations intrinsically programmed for survival after adoptive transfer and to modulate the immune status of recipients to prevent/delay antitransgene rejection responses.

Key Words: Cellular immunotherapy, Adoptive therapy, T lymphocyte, Clinical trial

 

Back to Article Outline

Introduction 

Although conventional chemotherapy, radiation therapy, and antibody therapy can be efficacious in treating lymphoma, relapse and progressive disease are the major sources of patient morbidity and mortality 1, 2. Experimental evidence that the cellular immune system can eradicate lymphoma provides a basis for the development of therapies aimed at amplifying antitumor immune responses 3, 4. The adoptive transfer of lymphoma-specific T cells is one strategy to augment antilymphoma immunity. A significant challenge to executing this strategy is the isolation of T cells specifically reactive to lymphoma. Alternately, the ex vivo derivation of tumor-specific T lymphocytes by genetic modification to express tumor-targeting chimeric antigen receptors (CARs) is a rapidly evolving focus of translational cancer immunotherapy 5, 6. Antibody-based CARs are HLA-unrestricted and thus can be used in patient populations with target-antigen–positive tumors.

We have constructed 2 CARs specific for the B cell lineage antigens CD20 and CD19 for the purpose of targeting lymphomas and leukemias 7, 8. When expressed in cytotoxic T lymphocytes (CTLs), these CARs redirect effector cells to lyse B-lineage lymphoma targets 7, 8. Here we report our initial clinical experience in manufacturing and infusing autologous T cells expressing CD20R or CD19R in patients with relapsed B cell lymphoma under City of Hope–held FDA-authorized trials BB-IND-8513/IRB 98142 and BB-IND-11411/IRB 01160, respectively.

Back to Article Outline

Materials and Methods 

Patients 

City of Hope Internal Review Board (IRB) protocols 98142 and 01160 were activated for patient accrual following IRB and Institutional Biological Safety Committee approval, FDA authorization (BB-IND-8513 and BB-IND-11411, respectively), and National Institutes of Health Office of Biotechnology Activities registration (9907-330 and 0207-543, respectively). In brief, for IRB 98142, patients were eligible if they had immunohistopathologically documented CD20+ diffuse large cell lymphoma (DLCL) with a history of recurrent or refractory disease and did not have central nervous system metastases. After leukapheresis, patients began salvage/mobilization chemotherapy, then underwent hematopoietic stem cell transplantation (HSCT). The first of 3 escalating-dose T cell infusions was given at 28 days post-HSCT. For IRB protocol 01160, patients were eligible if they had pathologically documented follicular lymphoma (FL) with evidence of progression after previous rituximab therapy and did not have central nervous system metastases or a history of allogeneic HSCT. These patients were enrolled no sooner than 3 weeks after their most recent cytotoxic chemotherapy.

Plasmid Vectors 

The plasmid expression vectors encoding the CD20R chimeric immunoreceptor and the neomycin phosphotransferase cDNAs and the CD19R chimeric immunoreceptor and the selection-suicide HyTK (a fusion of hygromycin resistance and HSV-1 thymidine kinase suicide genes) cDNAs have been described previously 7, 8 (Figure S1A). In brief, the chimeric construct consists of VH and VL gene segments of the CD20-specific Leu-16 or CD19-specific FMC63 monoclonal antibodies (mAbs), an IgG hinge-CH2-CH3 region, a CD4 transmembrane region, and the cytoplasmic domain of the CD3ζ chain (Figure S1B).

Isolation, Transfection, Selection, Cloning, and Expansion of T Cells 

The methods for OKT3 stimulation of peripheral blood mononuclear cells (PBMCs), and for PBMC electroporation, selection, cloning (IRB 98142 only), and subsequent growth using the rapid expansion method (REM), consisting of recursive 14-day cycles of activation with OKT3, recombinant human interleukin (rHuIL)-2, and PBMC/lymphoblastoid cell line (LCL)-irradiated feeders, have been described previously [9]. The overall T cell product manufacturing schemas for each trial are depicted in Figure S1C.

Cell Product Quality Control Tests 

The cell product quality control tests (QCTs) performed and the requisite test results for product release are summarized in Table S1.

Confirmation of Plasmid Vector Integration (IRB 98142 Only) 

A single site of plasmid vector chromosomal integration was confirmed by Southern blot analysis of XbaI/HindIII-digested T cell genomic DNA using a 420-bp NeoR-specific probe generated using the pcDNA3.1(-) plasmid as a template [9]. The pass criterion of this test was defined as detection of a single band.

Confirmation of CAR Expression 

Western blot analysis for CAR expression has been described previously [10]. In brief, reduced whole-cell lysates are subjected to Western blot analysis with an anti-human CD3-ζ (cytoplasmic tail)-specific mAb 8D3 (BD Pharmingen, San Diego, CA). This probe detects both the 16-kDa endogenous ζ and the 66-kDa CAR ζ. Pass criteria were defined as visualization of both the 16-kDa and 66-kDa bands. Flow cytometry analysis for surface CAR expression was determined using a fluorescein isothiocyanate (FITC)-conjugated Fc-specific antibody (Jackson ImmunoResearch, West Grove, PA). Pass criteria were defined as unimodal positive staining for Fc compared with the FITC-conjugated isotype control (BD Biosciences, San Jose, CA).

Surface Phenotype Determination 

T cell products were evaluated for cell-surface phenotype using standard staining and flow cytometric procedures with FITC-conjugated mAbs (BD Biosciences), followed by analysis on a FACScaliber analyzer (BD Biosciences). The pass criterion was ≥90% positive staining for TCR-αβ and CD8 (IRB 98142) or CD3 (IRB 01160) compared with the isotype control. Independent of the QCT guidelines, other correlative surface markers included CD4 for IRB 98142 and both CD4 and CD8 for IRB 01160.

Assay for Antilymphoma Cytolytic Activity 

Cytolytic activity of CAR+ CTLs against 51Cr-labeled human lymphoma Daudi cells was assessed as described previously using a 4-hour chromium release assay [8]. The pass criterion was ≥50% specific lysis at an effector-to-target ratio of 25:1.

Viability 

Viability was determined by standard trypan blue dye exclusion. The pass criterion was >90% viability.

Sensitivity to Ganciclovir Ablation (IRB 01160 Only) 

To test for acquired cytocidal sensitivity to ganciclovir (GCV), aliquots of cells were harvested from 5-day REM cultures, then maintained for 14 days in 37.5 U/mL rHuIL-2 with or without 1 μM GCV. Then the cells were harvested and subjected to viability testing. The pass criterion was ≤25% viability in the GCV-treated cultures.

Assay for Antigen/IL-2–Independent Growth 

First, 5 × 106 cells were washed and plated in antigen- and IL-2–free culture media at the end of a 14-day REM cycle. Parallel cultures of Jurkat T cells (American Type Culture Collection, Manassas, VA) (IRB 98142) or T cells cultured in the presence of 37.5 U/mL rHuIL-2 (IRB 01160) served as controls for expansion and viability. For IRB 98142, following an 11-day incubation, cultures were harvested, counted using trypan blue, plated into 96-well plates at 6000 viable cells per well, and pulsed with 1 μCi of 3H-TdR. DNA was harvested following a 4-hour incubation at 37°C. For IRB 01160, viable cell numbers of 14-day cultures were determined by flow cytometry as described previously [11]. Release criteria specified that cells must exhibit <10% of the Jurkat cpm (IRB 98142) or <10% of the IL-2+ control cell number (IRB 01160).

Sterility 

Sterility tests were performed according to an FDA Center for Biologics and Evaluation of Research–mandated schedule. Aliquots of media from the T cell cultures were plated onto bacterial and fungal growth media. Mycoplasma detection was conducted on media aliquots using the Gen-Probe Mycoplasma Tissue Culture-NI Rapid Detection System (Gen-Probe, San Diego, CA), and endotoxin levels were determined by enzyme-linked immunosorbent assay. Pass criteria were negative bacterial, fungal, and mycoplasma results, along with an endotoxin level <5 EU/kg recipient weight.

Adoptive Transfer of T Cells 

Processed and cryopreserved cell banks were thawed and expanded in culture to the desired cell numbers before being resuspended in 0.9% NaCl with 2% human serum albumin in a clinical reinfusion bag. T cells were reinfused i.v. over 30 minutes through either a central line or an age-appropriate sized i.v. catheter inserted into a peripheral vein. The infusion bag was mixed gently every 5 minutes during the infusion. The intrapatient dose escalation plan is shown schematically in Figure S2. In IRB 01160, fludarabine (Flu) was administered after the first T cell infusion as a potential nonmyeloablative (NMA) immunosuppressive regimen for attenuating possible rejection responses against the transferred T cells. The guidelines provided in the National Cancer Institute's (NCI) Common Toxicity Criteria, version 2.0 (https://ctep.ifo.nih.gov/l) were followed for the monitoring of toxicity and adverse event reporting. Rules for dose escalation, de-escalation, and cancellation were strictly enforced and resulted in 3 of the 4 treated patients deviating from the planned infusion cell dose escalation at least once.

In Vivo Persistence of Transferred T Cells 

For IRB 98142, PBMCs from heparinized peripheral blood samples were isolated and analyzed for percentage of transfected cells by quantitative polymerase chain reaction (qPCR) as described previously [12] using primers and probes to quantify CD20R copy number (details available on request).

For IRB 01160, samples were received, processed, stored, and analyzed in accordance with current good laboratory practice guidelines. A validated qPCR-based assay to quantify CD19R plasmid vector DNA in samples was developed and performed using an MJ Research DNA engine with a Chromo 4 continuous fluorescence detector qPCR module (Bio-Rad, Hercules, CA). Real-time qPCR was performed in a 20-μL reaction mixture volume containing 50 ng DNA, 10 μL of IQ SYBR Green Supermix (Bio-Rad; catalog no. 170-8880), and 0.5 pmol of each primer. Quantification of the CD19R transgene sequence in DNA isolated from patient PBMCs was evaluated using qPCR to amplify a 182-nucleotide fragment that spanned the CD4 transmembrane–zeta junction within the transgene coding sequence, and a standard curve derived by dilution of DNA isolated from a clone with a single integration of the CD19R transgene (primers, probes, and amplification conditions available on request). The qualification studies for this amplification reaction demonstrated no amplification from healthy donor-derived PBMCs (n = 5), whereas the transgene sequence could be quantified in a PBMC sample if the transgene containing DNA composed as little as 0.1% of the total DNA sample.

Analysis of Antitransgene Rejection Responses 

Pretreatment and post-treatment PBMCs (on days +75, +77, and +50 from UPN006, UPN009, and UPN035, respectively) were first stimulated with irradiated therapeutic T cells (3000 rads) or LCLs with and without pcDNA3.1(-) plasmid (8000 rads) plus irradiated pretreatment PBMCs as feeder cells at a 10:1 responder-to-stimulator ratio in RPMI 1640 (Irvine Scientific, Santa Ana, CA) supplemented with 2 mM L-glutamine (Irvine Scientific), 25 mM Hepes (Irvine Scientific), 100 U/mL penicillin, 0.1 mg/mL streptomycin (Irvine Scientific), and 10% heat-inactivated human serum. One week later, the same irradiated stimulator, as well as irradiated pretreatment PBMC feeders (3500 rads), were added at a 1:1:1 responder-to-stimulator-to-feeder ratio. This stimulation schema was repeated up to 2 more times (once weekly) until sufficient numbers were obtained for chromium-release assays. Cytolytic activity of these stimulated PBMC against 51Cr-labeled targets was analyzed as described previously using a 4-hour chromium-release assay [8].

For IRB 01160, cellular antitransgene immune responses were evaluated directly ex vivo using a combination of T cell receptor (TCR) Vβ spectratyping and CD107 degranulation assays. For the TCR Vβ spectratyping analysis, RNA was isolated from PBMC collected before and after infusion using the RNAqueous-4 PCR Kit for Isolation of DNA-free RNA (Applied Biosystems/Ambion, Austin, TX), and cDNA was then synthesized using the iScript cDNA Synthesis Kit (Bio-Rad). TCR Vβ spectratyping analysis was performed on cDNA samples essentially as described previously [13] using pools of Vβ-specific primers. A parallel series of amplifications using cDNA generated from pooled healthy donor PBMCs was performed as a quality control for amplification of each Vβ family. Aliquots of the amplification mixes were run on sequencing gels, followed by analyses using Genemapper v3.7 software (Applied Biosystems). The CD107 degranulation/mobilization assay was performed essentially as described previously [14], using patient PBMCs collected before and after infusion as effectors and infused T cell product or OKT3-expanded preinfusion PBMCs (i.e., autologous T cells) as targets. To detect spontaneous degranulation, a control sample without target cells was included in every experiment. FITC-conjugated anti-CD107a and anti-CD107b (BD Biosciences) were added directly to the tubes before incubation. After 5 hours of coincubation, cells were washed twice and stained with PE-Cy5-conjugated anti-CD8β and PE-conjugated anti-TCR Vβ23 (Beckman Coulter, Fullerton, CA) for 30 minutes at room temperature in the dark, rewashed, and analyzed on a FC500 flow cytometer using FCS Express v3.0 software (Beckman Coulter), with gating on CD8β+ and Vβ23+ lymphocytes.

Serologic Anti-CAR Immune Response Analysis 

Serum was isolated from patient blood samples collected in red-top (no additive) tubes using an established laboratory standard operating procedure and a qualified assay to detect CAR-specific serologic responses in samples. Samples were allowed to clot for 2-1/2 hours at room temperature, then centrifuged at 1000 × g for 15 minutes at 4°C. Serum was collected, aliquoted, and frozen immediately at −80°C. Flow cytometry detection of potential serum antibody responses against the anti-CD19R transgene was performed using parental versus CD19R-expressing Jurkat cell lines as indicator cell lines. The presence of antibodies in patient serum that specifically bound to CD19R+ Jurkat cells was evaluated by a subsequent incubation with FITC-conjugated AffiniPure F(ab')2 fragment goat anti-human IgG (Fcγ; Jackson ImmunoResearch). The cutoff for a negative response was established by defining the 95% one-sided prediction interval using a pool of non–CAR-reactive serum samples from healthy volunteers.

Back to Article Outline

Results 

Patient Characteristics 

The patients in these studies either had DLCL (BB-IND 8513/IRB 98142) or follicular non-Hodgkin lymphoma (NHL; BB-IND 11411/IRB 01160) (Table S2). One of the 4 research participants had bulky disease, including sites at the neck, chest, lymph nodes, and pelvis, at the time of enrollment. Three of the 4 participants had received rituximab (chimeric mAb specific for CD20) therapy before the first infusion of therapeutic T cells. The average duration of time from leukapheresis to first infusion was 106 days, and was affected by the time required to manufacture the T cell product and/or the timing of the patient's recovery from salvage therapy.

Generation of Genetically Modified T Cells 

Cell products meeting all quality control release tests (Table S1) were successfully generated for 2 of the 5 patients enrolled on IRB 98142, and for both patients enrolled on IRB 01160. The failure to release products for 3 of the patients enrolled on IRB 98142 stemmed from an inability to isolate T cell clones that expressed CD8, expressed endogenous TCR, or expanded adequately in vitro. The results of Southern blot analysis indicating the desired single-site insertions of the CD20R transgene within the released clones of IRB 98142 are depicted in Figure 1. Western blot and cell-surface expression profiles of T cell products for both trials are also depicted, confirming expression of the CAR protein. These cells were further subjected to flow cytometry analysis for confirmation of the T cell subset markers CD4, CD8, and either TCR-αβ or CD3. All of the cell products used in therapy also exhibited redirected killing of CD19+ and CD20+ human Daudi lymphoma targets in 4-hour chromium release assays. Furthermore, all of the cell lines retained their dependence on exogenous rHuIL-2 for survival and proliferation, and the HyTK-expressing lines of IRB 01160 tested positive for sensitivity to GCV-mediated ablation.

  • View full-size image.
  • Figure 1 

    T cell products meet release requirements. Depicted from left to right: Southern blots of T cell genomic DNA using an HyTK-specific probe showing existence of single bands as indicated by arrows; Western blots revealing both the 16-kDA endogenous CD3ζ and the 66-kDA CE7R chimeric ζ bands detected with anti-human CD3ζ cytoplasmic tail specific antibody; flow cytometry analysis for surface expression of the chimeric receptor using anti-Fc antibody, or for the T cell markers CD8, CD4, and TCR or CD3, where isotype control staining is indicated with the open histogram; ability of CTL clones to lyse CD19+ CD20+ Daudi targets was determined in a 4-hour 51Cr release assay; ganciclovir (GCV) sensitivity using a flow cytometry–based assay for viable cell numbers after 14 days of culture with either rHuIL-2 or rHuIL-2 + GCV; assays for IL-2 dependence were performed using 3H-thymidine incorporation measurements (cpm) of Jurkat T cells versus the indicated T cell clones after 11 days of culture in the absence of rHuIL-2 (UPN006, UPN009), or using a flow cytometry–based assay for viable cell numbers after the T cell products were cultured in the presence versus the absence of rHuIL-2 for 14 days (UPN035 and UPN037). N.D., not done.

Treatment Experience 

As depicted in Figure 2, the intrapatient dose escalations were carried out as planned (compare with Figure S2), with the exception that the 1010/m2 cell dose was never given in IRB 98142 because of protocol toxicity dose modification rules (UPN006 and UPN009). Indeed, because of grade 2 hepatic toxicities that were noticed with the first infusion dose of 108/m2 in UPN006, the second infusion was repeated at 108/m2, followed by an escalation to 109/m2 for the third infusion. Patient UPN009 exhibited a drop in hemoglobin after the second infusion that, although clinically insignificant (from 10.6 to 8.6), represented a Common Toxicity Criteria grade change from grade 1 to grade 2 anemia status, requiring repetition of the 109/m2 dose for the third infusion based on the protocol's defined rules for dose escalation. In UPN006, the second infusion was cancelled and rescheduled because of a puncture in the bag, which compromised the integrity of the T cell product. In UPN037, the last infusion was cancelled because of the detection of contaminated T cell product. Neither myeloablation (MA) followed by HSCT nor Flu resulted in a drop in absolute lymphocyte count (ALC) below the normal range (Table S3).

  • View full-size image.
  • Figure 2 

    Treatment regimens for each patient. First i.v. infusions of T cells were administered on day 0 for each patient. For UPN006 and UPN009, fractionated total body irradiation (TBI) and/or myeloablative chemotherapies administered to UPN006 and UPN009 are indicated just before administration of CD34+ autologous stem cells. BCNU, bis-chloronitrosourea; cytoxan, cyclophosphamide; VP-16, etoposide. For UPN035 and UPN037, administration of fludarabine (i.v. at 25 mg/m2) occurred between days 4 and 8 after the first T cell infusion, and rHuIL-2 administration (5 × 105 IU/ m2 BID) was initiated after the third T cell infusion.

There were no grade 3 or higher adverse events with a possible correlation to administration of 108 T cells per m2. However, examination of the adverse events at 109 T cells per m2 revealed one case of grade 3 self-limited lymphopenia in both IRB 98142 and IRB 01160, possibly attributed to cell administration (Table 1). At 2 × 109 T cells per m2, grade 3 lymphopenia and grade 3 eosinophilia each occurred once in IRB 01160; both resolved spontaneously with no adverse sequelae to the patients. Overall, the safety profile of this adoptive transfer therapy was acceptable.

Table 1. Adverse Event Summary
IRB TrialT Cell Dose, Cells/m2Event∗Occurrences

98142109Lymphopenia1
01160109Lymphopenia1
2 × 109Lymphopenia1
Eosinophilia1

IRB indicates internal review board; NCI, National Cancer Institute.

∗Only events of grade 3 or higher, according to the NCI Common Toxicity Criteria, with possible attribution to T cell administration are reported.

Follow-Up Clinical Status of Patients 

Although this was a phase I clinical trial with a primary purpose of determining safety, we also monitored the disease and survival status of each patient. For IRB 98142 (CD20R; DLCL), UPN006 (last infusion on February 24, 2000) relapsed in September 2001, whereas UPN009 (last infusion on November 22, 2000) continues to be in remission after autologous HSCT. At the time of the writing of this report, both UPN006 (after additional treatment) and UPN009 are alive and in remission. For IRB 01160 (CD19R; FL), UPN035 (last infusion on May 4, 2006) presented with a new diagnosis of CD19+/CD20+ DLCL in June 2006, and died in June 2007. UPN037 (last infusion on February 15, 2007) displayed progression on computed tomography scan in September 2007, and is currently alive and undergoing additional treatment.

In Vivo Persistence of Transferred T Cells 

Quantitative PCR performed to detect CD20R and CD19R plasmid copy numbers in PBMCs as a surrogate marker of the presence of adoptively transferred T cells showed varying T cell persistence among patients (Figure 3). Only 1 of the 4 patients (UPN006) had a detectable level of transferred T cells at 1 week after the first infusion of 108 cells/m2. A detectable level of transferred T cells at 1 week after infusion of 109 cells/m2 was found in only 2 of the 7 higher doses (UPN006 infusion 3 and UPN009 infusion 2), and at no time were transferred T cells detected at 1 week after infusion of 2 × 109 cells/m2. Thus, adoptively transferred T cell persistence did not appear to correlate with cell dose. Compared with the persistence after the initial infusion, UPN009, UPN035, and UPN037 also displayed significantly reduced levels of transferred T cells 24 hours after each additional infusion, suggesting the possibility of an antitransgene immune response mounted against the administered T cells (Figure 3).

  • View full-size image.
  • Figure 3 

    Transferred T cells do not persist long term in vivo. Using real time quantitative PCR, the percent of cells in the PBMC that were positive for the CD20R (A) or CD19R (B) genes were determined as an indicator of the relative amount of chimeric receptor expressing T cells in the PBMC samples collected at the indicated days during the treatment schedule. Escalating infusion doses are indicated by arrows. ∗Cells not harvested.

Detection of Transgene-Specific Immune Responses 

For IRB 98142 (CD20R; DLCL), the development of cellular immune responses against the infused T cell products was evaluated. For these analyses, PBMCs were collected from UPN006 and UPN009 before and after T cell administration, and, after in vitro stimulation, were compared for cytotoxic activity using chromium-release assays (Figure 4). The use of irradiated lymphoblastoid cells (xLCLs) to stimulate the PBMCs resulted in successful lysis of 51Cr-labeled LCLs, indicating that functional effector cells could be derived from each patient's PBMC samples. Interestingly, when the irradiated autologous T cell clones were used to stimulate the PBMCs, cytotoxic responses were seen against the 51Cr-labeled T cell clone only in the posttreatment sample collected from UPN009 (Figure 4A). This immunoreactivity against the T cell clone 6D10 used in therapy appeared to be specific for neomycin phosphotransferase but not the CAR, because cytotoxic responses could be observed against 51Cr-labeled LCLs that had been transduced with the pcDNA3.1(-) vector, which directs the expression of neomycin phosphotransferase, but lacks the CD20R transgene (Figure 4B). To better analyze the specificity of the rejection response, UPN009's posttreatment PBMCs that had been stimulated with irradiated 6D10 cells were cloned in limiting dilution; all clones were similarly specific for NeoR (Figure 4C).

  • View full-size image.
  • Figure 4 

    Transgene rejection response detected when T cells administered after HSCT. In the trial targeting CD20+ diffuse large-cell lymphoma, PBMCs collected before treatment and at day 75 (UPN006) or day 77 (UPN009) after initiation of treatment were stimulated in vitro with irradiated LCL as a control (xLCL) or the corresponding irradiated CTL clone that had been administered (i.e., x1A11 or x6D10). Effectors were then used in a 4-hour 51Cr-release assay using either LCL or the corresponding CTL as targets (A) or, in the case of UPN009, using LCL that had been transfected with the pcDNA3.1(-) vector lacking the CD20R transgene as targets (B). (C) Clones derived from UPN009 day 98 PBMC were also stimulated in vitro with irradiated 6D10 CTL and then analyzed for cytolytic activity against 51Cr-labeled LCL, 6D10, or pcDNA3.1(-) vector transfected LCL to determine specificity of transgene-specific response. Percent 51Cr-release at an E:T of 25:1 in each case is depicted for four representative clones.

For IRB 01160 (CD19R; FL), the development of both antibody and cellular immune responses against the infused T cell products was evaluated. For the antibody analyses, serum collected from UPN035 and UPN037 at enrollment and after T cell administration were both negative for antibody reactivity against the surface-expressed CD19R transgene using a flow cytometry–based assay (Figure 5A); however, evidence was found for cell-mediated immunoreactivity against the infused T cells. Examination of the TCR Vβ gene repertoire through spectratyping sequence analysis of PBMCs collected from UPN035 and UPN037 both before and after T cell administration revealed alterations in the Vβ profiles of the posttreatment PBMCs, with the appearance of unique clonotypes in the postinfusion samples indicative of a new immunoreactive response (Figure 5B). Furthermore, flow cytometry analysis of the TCR Vβ23+ and Vβ14+ subpopulations in UPN035's posttreatment PBMCs collected 2 weeks after the first infusion showed significant surface CD107 expression, an indicator of lysis-associated degranulation, on coculture with the infused T cell product (Figure 5C and data not shown). This specific degranulation was not observed with the pretreatment PBMCs or when the posttreatment PBMCs were cocultured with control T cells. Similar flow cytometric assays could not be carried out with UPN037's PBMC because of the lack of commercially available antibodies specific for the TCR Vβ genes (Vβ15 and Vβ21) that arose in this patient. The pretreatment and posttreatment PBMCs from UPN035 also were stimulated in vitro with irradiated LCLs or infused T cell product and compared for their cytotoxic activity using chromium-release assays (Figure 5D). As seen in UPN009 in IRB 98142, functional effector cells were derived from both of UPN035's PBMC samples, but cytolytic activity against the 51Cr-labeled T cells was seen only in the posttreatment sample. Taken together, these data suggest that, at least in some cases, the lack of T cell persistence observed in these 2 trials was because of immune rejection responses mounted by the patient's endogenous T cells.

  • View full-size image.
  • Figure 5 

    Rejection response detected when T cells were administered following fludarabine administration. (A) In the trial targeting CD19+ FL, serum collected at the time of patient enrollment (Pre-Trtmt) and at day 50 (UPN035) or day 42 (UPN037) after initiation of treatment was examined for immunoreactivity against Jurkat cells expressing the CD19R (red line) in a flow cytometry based assay. Parental Jurkat cells (grey histogram), and a known nonreactive serum (black line) were used as negative controls. (B) TCR Vβ profiles of the infused T cell product, day 0 PBMCs (collected just before first T cell infusion; Pre-Trtmt), and day 14 PBMC (collected before the second T cell infusion, Post-Trtmt) were determined by spectratyping analysis. Alterations in Vβ usage that were observed pretreatment versus posttreatment are highlighted by red boxes. (C) The TCR Vβ23+ population of pretreatment and day 14 PBMC from UPN035 was further analyzed by flow cytometry for surface CD107 expression as a marker of degranulation on coculture with the infused T cell product (infused T), or nonmodified autologous T cells (neg T). (D) Pretreatment and day 50 PBMC from UPN035 were stimulated in vitro with irradiated LCL as a control (xLCL) or with the irradiated T cell product (xTcells). Effectors were then used in a 4-hour 51Cr-release assay using either LCL or T cells as targets.

Back to Article Outline

Discussion 

More than 55,000 new cases of NHL are diagnosed each year in the United States, and the incidence of this disease is increasing 15, 16. Intermediate-grade B cell lymphomas (ie, diffuse large cell, mantle cell, marginal zone) and low-grade FLs are the most common subtypes of NHL, accounting for approximately 80% of cases. Most patients have widespread disease at the time of diagnosis and are treated with some combination of chemotherapy, radiation therapy, and rituximab. Unfortunately, more than two-thirds will relapse with their disease, and only 10% of these patients can be salvaged [17]. Efforts to improve survival in recurrent NHL focus primarily on the use of MA conditioning and autologous HSCT 18, 19, 20, 21, a strategy that is curative in approximately 46% of selected patients. However, the selected group of salvageable patients (aged <60 years, complete remission after primary treatment, and no known marrow or central nervous system disease) represents less than one-third of those with relapsed intermediate grade lymphomas. Patients with chemotherapy-resistant recurrent disease have a <15% 5-year event-free survival after HSCT, and those with refractory disease at the time of transplantation are rarely cured. Similarly, patients with mantle cell lymphoma and low-grade FL, whose disease becomes refractory to chemotherapy and radiation, have a poor prognosis despite high-intensity salvage therapy 22, 23, 24, 25. These findings have prompted the evaluation of additional strategies to eradicate lymphoma minimal residual disease after cytoreductive chemotherapy/radiation/rituximab, including the immunotherapeutic targeting of malignant B cells with adoptively transferred antigen-specific T cells.

Here, we have described our initial experience testing the feasibility and safety of lymphoma adoptive therapy with CTLs genetically modified to express redirecting CD20- and CD19-specific CARs. Our T cell production platform relied on plasmid vector electrotransfer into patient PBMC preparations. Although this approach facilitated regulatory approval and diminished expenses relative to the use of a viral vector platform, the low efficiency of chromosomal integration and sustained transgene expression encumbered the production platform to multiple rounds of activation/propagation in selection drugs (G418 and hygromycin B). Nevertheless, drug-resistant CAR+ T cells were isolated in each of the 7 enrolled patients. The reason why 3 of the enrolled subjects on protocol IRB 98142 did not have clones released was that only CD4+CD8- clones were isolated, when the release criteria specified CD4-CD8+ clones. This skewed result in the production runs was specific to lymphoma patients in this trial and likely reflects the repertoire changes in these patients because of disease and/or previous therapy at the time of apheresis for T cell production. Another observed limitation in the production platform during generation of polyclonal lines in IRB 01160 was the discordance between CAR expression and hygromycin resistance. The plasmid vector used in the trial drives the CAR and HyTK from 2 separate promoters, allowing chromosomal integration events that result in deletion of the CAR-encoding portion of the vector or transcriptional repression of the CAR promoter. As a result, we found demonstrable CAR expression in only a subset of polyclonal cell preparations. This problem potentially can be resolved in plasmid vectors using single promoter systems in which the two transgene open reading frames are separated by an internal ribosome entry site element or directly integrated into a single polypeptide with a cleavable linker element. Our group has now redesigned our platform to use self-inactivating lentiviral vectors and shortened ex vivo culture duration (∼28 days), improving the percentage of CAR-expressing T cells in polyclonal cell preparations and eliminating the need for bacterial drug-resistance gene coexpression.

The primary focus of these studies was to establish the safety of this approach. In this regard, T cell infusions were well tolerated up to 2 × 109 cells/m2. The most common event that could be attributed to T cell infusion was transient self-limited lymphopenia lasting less than 7 days. We suspect that this phenomenon is related to redistribution of the endogenous circulating repertoire as a consequence of infused cell product; whether it is based on cytokine/chemokine elaboration on activation or other mechanisms remains to be delineated. In the 2 patients who demonstrated immunologic rejection of the infused cell products, the third and subsequent cell doses in these 2 patients elicited a self-limited (<24 hours) febrile response with rigors. Despite the dramatic systemic febrile response, the patients did not exhibit cardiovascular instability or other overt toxicities associated with a “cytokine storm” syndrome. A toxic death proximal to cell infusion has been recently reported [26] in a patient with bulky chronic lymphocytic leukemia who received cyclophosphamide before administration of redirected T cells expressing a CD19-specific CAR with both CD28-costimulatory and CD3-ζ activation–signaling domains. The lack of serious toxicities in our patients might result from the limited numbers of circulating B cells at the time of T cell infusion; the IRB 98142 patients were 28 days from MA autologous HSCT, whereas the IRB 01160 patients experienced B cell reduction as a consequence of rituxan. However, analysis of peripheral blood samples after the last T cell infusion, as well as the follow-up clinical status of these patients, indicate that this strategy did not result in sustained B cell lymphopenia, as would be expected based on the transient engraftment of infused effector cells. Another possible explanation for the observed lack of toxicity is an attenuated cytokine response to activation based on the CAR having only a CD3-ζ–activation domain. Carefully designed future trials should test the effects of these parameters as they relate to the tolerability of cell infusions.

The NCI's Surgical Branch has demonstrated that the frequency and magnitude of melanoma-reactive tumor-infiltrating lymphocyte (TIL) engraftment can be enhanced by rendering patients lymphopenic before adoptive transfer and administering high-dose rHuIL-2 after transfer. TIL products were previously difficult to detect in very high numbers after adoptive therapy engraftment in about 50% of patients treated with the most intensive lymphodepleting regimens consisting of MA chemotherapy/TBI with CD34-selected stem cell rescue. Despite having received MA HSCT, the patients on IRB 98142 were not lymphopenic on day +28 when infusion of CD20-specific CD8+ clones commenced. Similarly, the patients on IRB 01160 were given a 5-day course of Flu at 25 mg/m2/dose without achieving lymphopenia before T cell transfer. Given the compelling data from the NCI melanoma trials, we plan to administer T cell products to lymphoma patients in conjunction with their autologous HSCT procedure on day +2, when lymphopenia is profound.

Our experience clearly identifies the issue of transgene immunogenicity as a mechanism that limits persistence. The plasmid electrotransfer platform required that we select drugs for stably integrated clones/lines. The rejection response observed was cellular and focused on NeoR in UPN009, whereas the transgene specificity of the rejection responses in UPN035 and UPN037 was not characterized further. The ability of transgenes expressed in T cells to elicit immune responses after adoptive transfer was clearly established by the findings of Berger et al. [27], in which the limited persistence of adoptively transferred HyTK+ T cells correlated with anti-HyTK transgene-specific immune responses. Furthermore, although studies by that group suggest that an NMA immunosuppressive regimen could prolong the in vivo persistence of T cells by attenuating possible rejection responses [28], Flu applied after the first dose of T cells in IRB 01160 apparently failed as an immunopreparative strategy to delete antitransgene-specific responses. Thus, more effective immunosuppressive/ lymphodepleting regimens might be advantageous for future clinical application. The use of more efficient vector transduction systems (eg, lentiviral vectors or the Sleeping Beauty system [29]) that would negate the requirement for ex vivo selection also might allow the omission of NeoR and HyTK. Even without the selection markers, the CARs themselves are expected to be immunogenic based on mouse single-chain variable fragments and fusion sites in the chimera. We expect that providing a window for cells to evade immunologic rejection by patient lymphodepletion will be the most practical strategy for limiting early rejection responses, whereas late rejection responses have the advantage of eliminating the gene-modified cells, which, if timed appropriately, could be exploited as a safety feature.

Poor in vivo persistence is the major problem in the cancer adoptive therapy field in general, likely related primarily to the intrinsic programming of T cells to survive after adoptive transfer. Effector T cells are inherently short-lived [30], and it has been suggested that acquisition of an effector phenotype during in vitro culture and expansion is a major reason for the poor survival of transferred T cells [31]. The adoptive transfer of virus-specific effector T cells from memory cell precursors can result in long-term repopulation in humans, however. Thus, the use of memory T cells, which are known to self-renew [32], and/or virus specific T cells, which would receive optimal costimulation after engagement of their native receptors, as populations for genetic redirection has become of increasing interest. Indeed, it has recently been reported that in neuroblastoma patients, Epstein-Barr virus-specific T cells engineered to coexpress tumor-specific receptors survived longer than those lacking virus specificity and were associated with tumor regression or necrosis in half of the subjects tested [33]. It also was recently reported that, in a macaque model of adoptive transfer, antigen-specific CD8+ T cell clones derived from central memory T (TCM) cells persisted long term in vivo, reacquiring phenotypic and functional properties of memory T cells and occupying TCM cell niches [34]. Accordingly, we have begun to develop a clinically compatible immunomagnetic selection system to isolate TCM cells from human PBMCs for subsequent processing to generate CAR redirected effector cells. We propose viral-specific, TCM-derived, anti–tumor effector cell infusion in lymphoma patients during profound lymphopenia shortly after autologous HSCT as the next logical iteration of our translational research in lymphoma immunotherapy.

Back to Article Outline

Acknowledgments 

This work was supported by the National Institutes of Health (Grants P01 CA30206 and P50 CA107399), the General Clinical Research Center (Grant M01 RR0004), the Lymphoma Research Foundation, the Marcus Foundation, and the Tim Nesvig Family Foundation. The authors thank Christine Wright, Araceli Hamlett, Cherrilyn Bautista, members of the City of Hope Center for Biomedicine and Genetics, and Dr. Shu Mi, and Dr. Ludmila Krymkaya, and Vivi Tran of the City of Hope Clinical Immunobiology Correlative Studies Laboratory for their technical assistance; and Merlita Alvarez, Lior Lewensztain, and Jamie Wagner for their help in compiling data.

Financial disclosure: Michael C. Jensen has a major ownership interest ($10,000 or more) as a patent holder. Laurence J. Cooper has minor ownership interest (<$10,000) as founder and majority owner of InCellerate Inc, a company that commercializes genetically modified T cells. None of the other authors has any conflicts of interest to disclose.

Back to Article Outline

Supplementary Data 

Download file

download text

Tables S1–S3

  • View full-size image.
  • Figure S1 

    Product manufacturing strategy. (A) Plasmid vectors used to genetically redirect the patient's T cells to recognize CD20 (left) and CD19 (right). CD20- and CD19-specific CAR sequences (CD20R and CD19R) are indicated, as well as promoter (CMVp, EF-1p, SV40p), poly adenylation (BGHpolyA, SV40polyA), drug resistance (AmpR, NeoR, Hy), and origin of replication (ColE1ori, F1 ori) sequences. Note that the HyTK sequence is a fusion of the hygromycin resistance gene, and the HSV-1 thymidine kinase suicide gene. (B) Schematic of the CD20- and CD19-specific CAR molecules. Each has a murine single-chain variable fragment (scFv) which makes it specific for either CD20 or CD19, a human IgG hinge-Fc (hugFc) domain, a human CD4 transmembrane (huCD4tm) domain, and a human CD3z cytoplasmic (huCD3zcyt) domain. (C) Product manufacturing schema of genetically modified T cells that target CD20 (left) and CD19 (right). Patient PSMCs were activated with the CD3 agonist OKT3 and rHuIL-2 for 3 days and then electroporated with linearized CD20R_pcDNA3.1(-) (left) or CD19R_HyTK-pMG (right). Cells were placed in drug selection conditions 2-3 days later. Cloning (for CD20R expressing products only) and expansion were carried out in the presence of OKT3, rHuIL-2, irradiated LCL, and irradiated PBMC feeders. T-25, T-75, and T-150 refer to flask sizes that permit 25, 50, and 100 mL of culture media respectively. At certain steps, the target number of selected CD20-specific T cell clones that were to be generated are indicated in the schema on the left. Performance of tests for viability, sterility, the presence of mycoplasma or endotoxin, transgene copy number (via Southern blot and PCR), total and surface CAR expression (via Western blot and flow cytometry), IL-2 dependence, and cytolytic activity against CD20- (left) or CD19- (right) expressing targets (micro-CRA, CRA) are indicated.

  • View full-size image.
  • Figure S2 

    Protocol treatment schema. (A) Treatment schema for recurrent/refractory CD20+ lymphoma using CD20R+ autologous T cells. Autologous HSCT was to be performed after a myeloablative preparative regimen of high dose chemotherapy with or without fractionated TBI; administration of escalating T cell dose infusions at 2-week intervals would then begin as early as 28 days later. (B) Treatment schema for CD19+ FL using CD19R+ HyTK+ autologous T cells. Lymphodepleting fludarabine was to be administered i.v. at 25 mg/m2 for 5 days; T cell supportive rHuIL-2 was to be administered s.c. at 5 × 105 IU/ m2 twice daily for up to 5 days after infusions 3 and 4.

Back to Article Outline

References 

  1. Kofler DM, Mayr C, Wendtner CM. Current status of immunotherapy in B cell malignancies. Curr Drug Targets. 2006;7:1371–1374
  2. Molina A. A decade of rituximab: improving survival outcomes in non-Hodgkin's lymphoma. Annu Rev Med. 2008;59:237–250
  3. Cesco-Gaspere M, Morris E, Stauss HJ. Immunomodulation in the treatment of haematological malignancies. Clin Exp Med. 2009;9:81–92
  4. Timmerman JM. Immunotherapy for lymphomas. Int J Hematol. 2003;77:444–455
  5. Sadelain M, Brentjens R, Riviere I. The promise and potential pitfalls of chimeric antigen receptors. Curr Opin Immunol. 2009;21:215–223
  6. Riddell SR. Engineering antitumor immunity by T-cell adoptive immunotherapy. Hematol Am Soc Hematol Educ Progr. 2007;250–256
  7. Jensen M, Tan G, Forman S, et al. CD20 is a molecular target for scFvFc:zeta receptor redirected T cells: implications for cellular immunotherapy of CD20+ malignancy. Biol Blood Marrow Transplant. 1998;4:75–83
  8. Cooper LJ, Topp MS, Serrano LM, et al. T-cell clones can be rendered specific for CD19: toward the selective augmentation of the graft-versus-B-lineage leukemia effect. Blood. 2003;101:1637–1644
  9. Jensen MC, Clarke P, Tan G, et al. Human T lymphocyte genetic modification with naked DNA. J Mol Ther. 2000;1:49–55
  10. Gonzalez S, Naranjo A, Serrano LM, et al. Genetic engineering of cytolytic T lymphocytes for adoptive T-cell therapy of neuroblastoma. J Gene Med. 2004;6:704–711
  11. Cooper LJ, Ausubel L, Gutierrez M, et al. Manufacturing of gene-modified cytotoxic T lymphocytes for autologous cellular therapy for lymphoma. Cytotherapy. 2006;8:105–117
  12. Park JR, Digiusto DL, Slovak M, et al. Adoptive transfer of chimeric antigen receptor re-directed cytolytic T lymphocyte clones in patients with neuroblastoma. Mol Ther. 2007;15:825–833
  13. Akatsuka Y, Martin EG, Madonik A, et al. Rapid screening of T-cell receptor (TCR) variable gene usage by multiplex PCR: application for assessment of clonal composition. Tissue Antigens. 1999;53:122–134
  14. Betts MR, Brenchley JM, Price DA, et al. Sensitive and viable identification of antigen-specific CD8+ T cells by a flow cytometric assay for degranulation. J Immunol Methods. 2003;281:65–78
  15. Alexander DD, Mink PJ, Adami HO, et al. The non-Hodgkin lymphomas: a review of the epidemiologic literature. Int J Cancer. 2007;120(Suppl 12):1–39
  16. Rogers BB. Overview of non-Hodgkin's lymphoma. Semin Oncol Nurs. 2006;22:67–72
  17. Arora NK, Hamilton AS, Potosky AL, et al. Population-based survivorship research using cancer registries: a study of non-Hodgkin's lymphoma survivors. J Cancer Surviv. 2007;1:49–63
  18. Gisselbrecht C, Bethge W, Duarte RF, et al. Current status and future perspectives for yttrium-90 ((90)Y)-ibritumomab tiuxetan in stem cell transplantation for non-Hodgkin's lymphoma. Bone Marrow Transplant. 2007;40:1007–1017
  19. Paolo C, Lucia F, Anna D. Hematopoietic stem cell transplantation in peripheral T-cell lymphomas. Leuk Lymphoma. 2007;48:1496–1501
  20. Santos ES, Kharfan-Dabaja MA, Ayala E, et al. Current results and future applications of radioimmunotherapy management of non-Hodgkin's lymphoma. Leuk Lymphoma. 2006;47:2453–247
  21. Zinzani PL. Autologous hematopoietic stem cell transplantation in non-Hodgkin's lymphomas. Acta Haematol. 2005;114:255–259
  22. Bertoni F, Zucca E, Cavalli F. Mantle cell lymphoma. Curr Opin Hematol. 2004;11:411–418
  23. Brody J, Advani R. Treatment of mantle cell lymphoma: current approach and future directions. Crit Rev Oncol Hematol. 2006;58:257–265
  24. Goy A, Feldman T. Expanding therapeutic options in mantle cell lymphoma. Clin Lymphoma Myeloma. 2007;7(Suppl 5):S184–S191
  25. Smith MR. Mantle cell lymphoma: advances in biology and therapy. Curr Opin Hematol. 2008;15:415–421
  26. Brentjens RJ, Riviere I, Hollyman D, et al. Unexpected toxicity of cyclophosphamide followed by adoptively transferred CD19-targeted T cells in a patient with bulky CLL. Mol Ther. 2009;17:S157;[Abstract]
  27. Berger C, Flowers ME, Warren EH, et al. Analysis of transgene-specific immune responses that limit the in vivo persistence of adoptively transferred HSV-TK–modified donor T cells after allogeneic hematopoietic cell transplantation. Blood. 2006;107:2294–2302
  28. Berger C, Huang ML, Gough M, et al. Nonmyeloablative immunosuppressive regimen prolongs in vivo persistence of gene-modified autologous T cells in a nonhuman primate model. J Virol. 2001;75:799–808
  29. Singh H, Manuri PR, Olivares S, et al. Redirecting specificity of T-cell populations for CD19 using the Sleeping Beauty system. Cancer Res. 2008;68:2961–2971
  30. Wherry EJ, Teichgraber V, Becker TC, et al. Lineage relationship and protective immunity of memory CD8 T cell subsets. Nat Immunol. 2003;4:225–234
  31. Gattinoni L, Powell DJ, Rosenberg SA, et al. Adoptive immunotherapy for cancer: building on success. Nat Rev Immunol. 2006;6:383–393
  32. Fearon DT, Manders P, Wagner SD. Arrested differentiation, the self-renewing memory lymphocyte, and vaccination. Science. 2001;293:248–250
  33. Pule MA, Savoldo B, Myers GD, et al. Virus-specific T cells engineered to coexpress tumor-specific receptors: persistence and antitumor activity in individuals with neuroblastoma. Nat Med. 2008;14:1264–1270
  34. Berger C, Jensen MC, Lansdorp PM, et al. Adoptive transfer of effector CD8 T cells derived from central memory cells establishes persistent T cell memory in primates. J Clin Invest. 2008;118:294–305

 Laurence J. Cooper is currently at the Division of Pediatrics, M.D. Anderson Cancer Center, Houston, Texas. Michael Kalos is currently at the Abramson Family Cancer Research Institute, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania.

 Financial disclosure: See Acknowledgments, page 1256.

PII: S1083-8791(10)00119-9

doi:10.1016/j.bbmt.2010.03.014

Biology of Blood and Marrow Transplantation
Volume 16, Issue 9 , Pages 1245-1256, September 2010

Article Tools

* Image Usage & Resolution