A Phase I study of 11 pediatric patients with newly diagnosed, Stage 4 neuroblastoma was conducted using monocyte-derived dendritic cells (DC) pulsed with tumor RNA to produce antitumor vaccines (DCRNA).
A Phase I study of 11 pediatric patients with newly diagnosed, Stage 4 neuroblastoma was conducted using monocyte-derived dendritic cells (DC) pulsed with tumor RNA to produce antitumor vaccines (DCRNA).
Patients received two courses of induction with carboplatin followed by standard chemotherapy, surgery, radiation, high-dose therapy, stem cell rescue, and DCRNA vaccine therapy.
The results showed that this method for producing and administering DCRNA from a single leukapheresis product was both feasible and safe in this pediatric neuroblastoma population. Two courses of carboplatin maintained lymphocyte counts at normal levels. However, immune function 6 weeks after high-dose chemotherapy and stem cell rescue and prior to receiving DCRNA was impaired in all patients tested. There was an alteration in the ratio of CD4-positive and CD80-positive T cells. CD4-positive cell numbers were below normal, whereas CD8-positive cell numbers were above normal for all patients. In addition, CD19-positive cell numbers were below normal for all but one patient. It was found that humoral responses to recall antigens (diphtheria and tetanus) and cellular responses to mitogen and recall antigens were below normal in most patients. Despite this, two of three patients tested showed a tumor-specific humoral immune response to DCRNA. Among the patients who had measurable disease at the time of DCRNA vaccine, none showed any objective tumor response.
DCRNA vaccines were both safe and feasible in children with Stage 4 neuroblastoma. Humoral responses to tumor were detected, although remained immunosuppressed at the time of administration, limiting efficacy. Cancer 2005. © 2005 American Cancer Society.
Neuroblastoma is the most common extracranial solid tumor found in children.1 The prognosis is good in children age < 1 year with localized disease; however, among older children with advanced disease, only 30% of patients are progression free 3 years after treatment.1 Current therapies include high-dose chemotherapy, radiotherapy, surgery, and stem cell support but even with these aggressive therapies, there is limited success reported for patients with advanced stage disease. There is an urgent need for new therapeutic strategies for advanced stage neuroblastoma. Immunotherapy may be an attractive alternative for treating children with neuroblastoma. In adult populations, it has been shown that dendritic cell (DC)-based antitumor vaccines are safe and feasible.2 DCs were pulsed with tumor lysate or with keyhole limpet hemocyanin and were mixed together to constitute antitumor vaccines for patients with Stage IV malignancies. That study showed that 7 of 14 patients had increased interferon γ (IFN-γ) production in response to tumor antigen.3 Pediatric studies using DC antitumor vaccines have been limited to date but may show promise with more study.4, 5 In recent years, there have been efforts to improve patient outcomes by incorporating nonchemotherapeutic adjuvant therapies in addition to standard chemotherapy regimens with varied success. For example, retinoids have proved to be an effective therapy for increasing event-free survival in patients with high-risk neuroblastoma after a standard course of chemoradiotherapy.6, 7
Other adjuvant therapies have included immunotherapy approaches, such as antibody therapies. It has been shown that anti-GD2 antibodies given with granulocyte-macrophage–colony stimulating factor are effective in preventing disease recurrence in patients who achieved remission after primary chemotherapy but were ineffective in children with progressive disease.8 One study using a bispecific antibody for neural cell adhesion molecule and CD3 stimulated both CD4-positive cells and CD8-positive cells and produced memory cells against neuroblastoma cells, hence demonstrating that antibody immunotherapies may be effective in neuroblastoma.9
The status of the immune system is of critical importance with respect to anticancer immunotherapy. Recent studies have focused on immunocompetency after chemotherapy, but baseline data prior to chemotherapy are lacking. Immune function was examined in patients with acute lymphoblastic leukemia, Hodgkin disease, or solid tumors after successful chemotherapy with or without radiotherapy.10 Immune responses to specific antigens were lower than normal for both the humoral and cellular arms of the immune system in most patients, with only 19% of patients demonstrating normal responses 12 months after chemotherapy.10 Others showed that, immediately after high-dose chemotherapy, humoral responses were quantitatively below protective levels for standard vaccines, such as tetanus and diphtheria; however, at 6 months after chemotherapy, most humoral factors had recovered. However, that study did not examine cellular immune function.11 Further studies involving the cytokine profiles of cancer patients showed that interleukin 2 (IL-2) is deficient in peripheral blood stem cells after traditional chemotherapy and bone marrow transplantation,12 and mononuclear cells in patients with advanced cancer show deficiencies in T-helper 1 responses (decreased IFN-γ, IL-10, and IL-12 and increased IL-4).13 Several studies have shown that, although other cellular blood components can recover relatively quickly after chemotherapy, lymphocytes, and CD4-positive T cells in particular, do not.14–17 However, because thymic function is an important pathway for CD4-positive reconstitution and becomes inactive with age, children have better CD4-positive T-cell recovery after chemotherapy compared with young adults.18 Despite these informative quantitative studies, to our knowledge qualitative immune function in children undergoing standard treatments for malignancies has not been studied systematically to date.
The current Phase I clinical study (also) provides important information on both the immunocompetency of patients with neuroblastoma and the efficacy of DCRNA vaccines to stimulate an immune response. The results of the current study suggest that, to optimize vaccine therapy in this type of cohort, future examination of the effects of cancer and its therapies on the immune system are warranted.
Protocols were reviewed and approved by the Ethics in Human Research Committee at The Royal Children's Hospital, Parkville, Victoria, Australia. Informed consent was obtained for all patients prior to enrolment. Eligible patients had untreated, advanced stage neuroblastoma and were either < 365 days and International Neuroblastoma Staging System (INSS) Stage 4 or n-myc-amplified Stage 4S, or > 365 days and Stage 4 or n-myc-amplified Stage 3. A source of tumor was mandatory to obtain material for histologic diagnosis, cytogenetics, molecular genetics, and RNA extraction. Standard staging evaluation included computed tomography or magnetic resonance imaging (MRI) scans with contrast, chest X-ray, Technetium-99m bone scan, metaiodobenzyl-guanidine scintigraphy (if available), bilateral bone marrow aspirates, and trephines. Patients were required to have adequate performance status (Karnofsky score > 50% or Lansky play performance scale > 50), and organ function.
Figure 1 shows that induction chemotherapy with leukapheresis commenced with 2 courses of carboplatin at a dose of 560 mg/m2 followed by granulocyte-colony stimulating factor (G-CSF)-mobilized leukapheresis if bone marrow clearing occurred. Carboplatin was used as a single agent because it has been demonstrated that it is well tolerated and effective, with a 77% response rate.19 Apheresis product was subject to reverse transcriptase-polymerase chain reaction analysis for neuroblastoma markers for tyrosine hydroxylase20 and PGP9.521 quality control, such that if the level of tumor cells in the collection was > 1 in 1000 cells, then it was discarded, and a repeat collection at a later time point was required. Apheresis was conducted to obtain aliquots for vaccine preparation (≥ 1 × 1010 mononuclear cells/m2) and stem cell rescue (≥ 2.5 × 106 mononuclear cells/m2). Courses 3, 4, and 6 consisted of doxorubicin at a dose of 60 mg/m2 on Day 1; cyclophosphamide at a dose of 2 g/m2 on Days 1 and 2; and vincristine at a dose of 1.5 mg/m2 on Days 1, 8, and 15. Course 5 consisted of cisplatin at a dose of 40 mg/m2 on Days 1–5 and etoposide at a dose of 200 mg/m2 on Days 2, 3, and 4. Induction chemotherapy courses were scheduled 3 weeks apart.
Postinduction surgery for residual primary disease was performed after six courses of chemotherapy, when possible, on all patients who achieved a measurable response to chemotherapy and had negative bone marrow. The objective of surgery was to document regional disease status, to achieve a complete response if resection was possible, or to reduce bulk tumor to minimal residual status.
Radiation therapy was performed immediately prior to intensive chemotherapy and stem cell rescue (Course 7). Radiation therapy was used to treat the primary tumor bed to a total dose of 24 grays (Gy) in 16 fractions or 30 Gy in 20 fractions for patients age < 36 months and age > 36 months, respectively. Radiation therapy was calculated based on diagnostic imaging performed after the completion of induction chemotherapy. For patients who achieved a complete response because of postinduction surgery, the treatment volume was the extent of disease based on postchemotherapy imaging plus a 2.0-cm margin.
Intensive chemotherapy was given with stem cell rescue. Course 7 included carboplatin at a calculated daily dose of 3 × uncorrected glomerular filtration rate (GFR) + (15 × surface area) mg to achieve an area under the concentration time curve of 3 mg/mL per minute and etoposide at a dose of 6.7 mg/kg per day, both given as a 96-hour continuous infusion on Days 6, 5, 4, and 3, as well as melphalan at a dose of 2.3 mg/kg per day on Days 6, 5, and 4. Stem cells were reinfused on Day 0, and G-CSF therapy was initiated on Day 1.
DCRNA vaccine was administered in the following manner: After a 6-week rest period, a combined diphtheria and tetanus (DT) vaccine was administered to each patient 1 week prior to the first DC vaccine to assess recall immunity to known antigens. DCRNA vaccines were administered to patients both intradermally (ID) and intravenously (IV) in Weeks 0, 2, and 4. Patients with stable disease were eligible for 3 subsequent vaccinations at 3-month intervals if treatments were well tolerated and if there was sufficient quantity of vaccine. ID vaccine was delivered using a 25-gauge needle on a 1-mL syringe, and the site of administration was rotated for each vaccination. A 3-tiered dose-escalation strategy using 3-patient cohorts was planned for the intravenous route, with low, medium, and high doses of 0.5 × 107 cells/m2, 1.5 × 107 cells/m2, and 5.0 × 107 cells/m2, respectively. The ID dose was 0.5 × 107 cells/m2. Patients were observed and vital signs were monitored before, during, and for 30 minutes after each vaccine was given, with a 1-hour interval between the 2 routes of administration. Toxicity was monitored according to The National Cancer Institute Common Toxicity Criteria (version 2.0). Autoimmune serologies were followed to detect any subclinical evidence of immune dysregulation. Immunologic, clinical, and MRI evaluations were performed at baseline, before and after the first three vaccinations, after each subsequent vaccination, and 1 month after the final vaccine. Standard response criteria for tumor response were used.22, 23
Tumor response was evaluated as follows: A complete response was defined as no evidence of primary or metastatic disease and normal urinary catecholamines. A very good partial response was defined as a reduction > 90% in tumor volume, healing of bone lesions, no evidence of tumor elsewhere, and normal urinary catecholamines. A partial clinical response was defined as a reduction > 50% in all measurable tumor volumes, no more than minimal bone marrow infiltration (only 1 sample positive for tumor cells and reduced from previous samples), and improvement in bony lesions. A minor response was defined as a 25–50% reduction in all tumor volumes and no new lesions. No response was defined as a decrease < 25% in measurable tumor volumes and no new lesions. Progressive disease was defined as an increase > 25% in the size of ≥ 1 lesions or the appearance of new lesions.
Tumor tissue was frozen at − 80 °C immediately after surgical removal at diagnosis. Extraction of total RNA was performed using RNeasy Maxi columns, and the optional DNAse treatment of the RNA column was performed using the RNase-free DNase set according to the manufacturer's instructions (Qiagen GmbH, Hilden, Germany).4
A peripheral blood monocyte count ≥ 100 cells/mm3 was considered sufficient to proceed with leukapheresis, which was performed as described by Caruso et al.4
For generation of clinical grade DC, methods previously described by Romani et al.24 were utilized with modifications described by Heiser et al.25 Briefly, processing of blood through restricted peripheral blood leukapheresis was performed for each patient, and the product was received in the laboratory at room temperature. Cells were diluted 1:1 with sterile saline and were purified further using density-gradient centrifugation over Ficoll-Hypaque plus reagent (Pharmacia Biotech, Uppsala, Sweden), and cells were resuspended at 6.5 × 106 cells/mL in serum-free AIM-V medium (Invitrogen, Carlsbad, CA). After 2 hours of humidified incubation at 37 °C, nonadherent cells were removed and cryopreserved for later assays. Adherent cells were cultured at 37 °C in serum-free AIM-V medium containing human recombinant IL-4 at 25 ng/mL (R&D Systems, Minneapolis, MN) and human recombinant granulocyte-macrophage–colony-stimulating factor at 800 ng/mL (BD PharMingen, San Diego, CA). After 7 days of culture, nonadherent cells were phenotyped by fluorescent-activated cell sorting analysis using fluorochrome-conjugated monoclonal antibodies to CD14-PE, human leukocyte antigen-D related (HLA-DR)-PC5, and CD11c-fluorescein isothiocyanate (Immunotec, Montreal, Quebec, Canada). Cells were considered suitable for vaccine use if they were > 50% HLA-DR-positive and < 15% CD14-positive.24
Pulsing of autologous DCs with RNA was performed by simple coincubation25, 26 in which 5 × 106 cells/mL in AIM-V medium (Invitrogen) were coincubated with 25 μg/mL tumor RNA at 37 °C for 45 minutes. DCRNA was washed twice with saline (Baxter, Deerfield, IL) and resuspended in freezing medium containing 80% autologous plasma, 10% saline, and 10% Cryoserv® (Edwards Lifesciences, Irvine, CA). Aliquots of 10–20 × 106 viable cells/mL were frozen in a rate-control freezer (Planer Kryolo series III).
Lymphocyte subsets were determined using the Simultest™ IMK lymphocyte kit (Becton Dickinson, San Jose, CA) as a clinical test, which was performed by the Immunology Laboratory at the Royal Children's Hospital. Briefly, peripheral blood samples were stored at room temperature for no longer than 24 hours before they were assayed. Next, 100 μL of whole blood were used for each test and were incubated with fluorochrome-conjugated antibodies against CD19, CD4, CD8, or CD16 plus CD56 for 30 minutes in the dark at room temperature. After lysing the red blood cells with lyse solution (0.8% NH4Cl buffer) for 10 minutes in the dark, samples were centrifuged for 5 minutes and were washed once with phosphate-buffered saline (PBS). Samples were then analyzed by flow cytometry on a Becton Dickinson FacSCAN™ (Becton Dickinson).
Mitogen-induced cellular proliferation was assessed by standard 3H-thymidine uptake, as described previously.27 Briefly, peripheral blood mononuclear cells (PBMCs) (1 × 105 per well) were cultured in AIM-V media supplemented with β-mercaptoethanol (5 × 10− 5 M; Sigma Chemical Company, St. Louis, MO) in a 96-well plate for 72 hours with (stimulated) or without (unstimulated) phytohemagglutinin (PHA) (30 μg/mL final concentration). Two hours prior to harvesting, 1 μCi per well 3H-thymidine (Amersham, Buckinghamshire, U.K.) was added to each well. Cells were harvested at 72 hours, and radioactivity was measured in a scintillation counter (TopCount-NXT; Packard Biosciences, Meriden, CT). Results are expressed as an average counts per minute (CPM).
Immunoglobulin G (IgG), IgA, and IgM antibody classes were analyzed at enrolment to determine whether this cohort of patients had normal levels of antibodies prior to vaccination with DCRNA. Patient serum samples were stored at 4 °C until IgG, IgA, and IgM totals were measured by standard nephelometry (Beckman Array; Beckman Coulter, Hialeah, FL) performed as a routine test by the Immunology Laboratory at the Royal Children's Hospital. Normal ranges were established as described previously.28
Specific IgG antibodies against tetanus and diphtheria were determined by enzyme-linked immunoadsorbent assay (ELISA) for each antigen as described by Caruso et al.4
Microtiter plates (Maxisorp™ plates; Nunc A/S, Roskilde, Denmark) were precoated overnight at 4 °C with 100 μg/mL tumor lysate or 100 μg/mL autologous PBMC lysate as control material in separate wells. Plates were then incubated with 1% bovine serum albumen/1% casein solution in PBS (blocking buffer) at 37 °C for 1 hour. After 1 wash with PBS, patient serum (diluted 1/50 in blocking buffer) was added to each well and incubated at 4 °C for 1 hour. Plates were washed 3 times with PBS and vortexed on a microplate vortex (Rohm Pharma) for 15 minutes at 900 revolutions per minute (rpm). Wells were incubated with Goat antihuman-Ig conjugated to horseradish peroxidase (Serotech, Toronto, Ontario, Canada) at 4 °C for 90 minutes and washed 3 times with PBS containing 0.5% Tween-20 (wash buffer). The plates were vortexed with wash buffer twice for 15 minutes at 900 rpm, washed twice for 5 minutes, then rinsed 3 times with deionized H2O. Antibody complexes were detected with 3,3′,5,5;-tetramethylbenzidine (Zymed Laboratories, San Francisco, CA) at room temperature for 20 minutes protected from light. Reactions were stopped using 2 N H2SO4, and the outer dimension was measured at 450 nanometers (nm) using a microplate reader (Multiskan Ascent; Labsystems, Helsinki, Finland). Positive and negative controls were comprised of pooled pediatric serum samples that were screened previously for reactivity to IMR-32 lysates. Samples were assayed in triplicate, and the results are reported as the average ± standard deviation.
The study sample size was determined according to traditional Phase I dose-level design. Student t tests for unpaired data were performed to obtain P values to determine significance. Error bars were generated to represent the standard deviation of the average of triplicate samples.
Patient characteristics are summarized in Table 1. In total, 11 patients (6 males and 5 females) ages 1.7–5.8 years are listed in order of enrollment. MYCN amplification was present in 4 of 11 patients. All treatment for each patient is shown in the order in which it was received. Patients 5 and 6 had early progressive disease. Patient 10 had unresectable primary disease, achieved only a minor response to 6 cycles of chemotherapy, and received no further therapy. Patient 11 was deemed to have unresectable primary disease but did complete all chemotherapy and radiation therapy and achieved a partial response.
|Patient no.||Age (yrs)||Gender||Primary site||Distant metastases||MYCN||Protocol treatment summarya||Responseb||No. of vaccines givenc||Toxicity||PFS (mos)||Current status||Length of follow-up (mos)|
|1||4.3||Male||R adrenal||BM||Amplified||Chemo (6 cycles), surgery, XRT HD chemo, and autoPBSCR||CR||4||Grade 1 skind||13||Dead||20|
|2||3.9||Female||L adrenal||Cervical LN||Not amplified||Chemo (6 cycles), surgery, XRT, HD chemo, and autoPBSCR||CR||3||None||17||Dead||19|
|3||5.0||Male||Mid-retroperitoneal||BM, orbit||Not amplified||Chemo (6 cycles), surgery, XRT, HD chemo, and autoPBSCR||VGPR||3||None||23||Dead||36|
|4||3.4||Male||R adrenal||BM, bone, orbit||Amplified||Surgery, chemo (6 cycles), XRT, HD chemo, and autoPBSCR||CR||4||None||16||Dead||21|
|5||1.7||Female||R adrenal, paraspinal/epidural||BM, bone||Not amplified||Chemo (4 cycles)||PD||0||NA||3||Dead||5|
|6||4.7||Female||L adrenal, retroperitoneal||BM, bone||Amplified||Chemo (6 cycles), surgery, XRT,||PD||0||NA||8||Dead||9|
|7||3.9||Male||L adrenal||BM, bone||Not amplified||Chemo (6 cycles), surgery, XRT, HD chemo, and autoPBSCR||PR||1||None||17||Dead||22|
|8||1.6||Male||L adrenal||BM||Amplified||Chemo (6 cycles), surgery, XRT, HD chemo, and autoPBSCR||CR||0||NA||10||Dead||13|
|9||5.8||Female||L retroperitoneal||BM, bone||Not amplified||Chemo (6 cycles), surgery, XRT, HD chemo, and autoPBSCR||VGPR||3||None||15||Dead||21|
|10||4.2||Male||R adrenal, retroperitoneal||BM, lung, liver||Not amplified||Chemo (6 cycles), unresectable primary||MR||0||NA||7||Dead||8|
|11||3.5||Female||Mid-retroperitoneal||BM, bone||Not amplified||Chemo (6 cycles), XRT, HD chemo, and autoPBSCR; unresectable primary||PR||4e||None||14||Alive||14|
Three patients were withdrawn prior to vaccine therapy due to progressive disease (Patients 5 and 6) or chemoresistant disease (Patient 10). Patient 8 did not receive DCRNA vaccines due to inadequate DC numbers. Seven patients went on to receive DCRNA vaccines and were evaluated for toxicity, feasibility, immune status, and antitumor immune responses.
RNA and DC production feasibility is shown in Table 2. This study was designed to test the use of a single leukapheresis preparation. Tumor RNA extraction was adequate for vaccine preparation in all patients. An average of 11.6 × 107/m2 DCRNA was prepared with 8 of 9 leukapheresed patients satisfying feasibility criteria (required at least 3 × 107/m2 DCRNA). The in vitro production of DCs for Patient 8 failed phenotypic standards for use due to lack of HLA-DR-positive cells and a higher than acceptable CD14-positive cell population.
|Patient no.||RNA (μg)||RNA (μg/g tumor)||PB monocytes (× 106/L)a||PBMNC (× 106)||PBMNC/m2 (× 106)||DCb|
|Prepulse PBMNC (%)||Prepared DCRNA/m2 (× 107)|
In addition, a 3-tiered dose escalation strategy using three patient cohorts, according to the order of enrolment, was planned for the IV route, with low, medium, and high doses of 0.5 × 107/m2, 1.5 × 107/m2, and 5.0 × 108/m2, respectively. Table 2 shows DCRNA adjusted for patient body surface area. Patients are listed in the order of enrolment, and all patients except Patient 2 had sufficient yields of vaccines. Although the feasibility criteria were met in eight of nine apheresed patients, dose escalation was not achievable to provide all six dose-escalated doses at anything above the low-dose level. Further escalation would have required multiple PBMC collections, which were not included in the study protocol. Therefore, all patients who received DCRNA vaccines were treated at the low dose.
Vaccinations were administered safely with no measurable toxicity, and none of the patients with standard vaccine preparation developed any clinical or biochemical signs of autoimmune disease (Table 1). However, the DCRNA vaccine received by Patient 1 included the addition of hepatitis B surface antigen to serve as an adjuvant for DCRNA and showed local inflammation at the injection site subsequent to the initial vaccine administration. Due to the very high cell death that the addition of hepatitis B surface antigen caused to the DCRNA vaccine preparation, we chose to remove this step from the study protocol for subsequent patients.
Four of seven patients had measurable disease at the time of DCRNA vaccine. None of these patients demonstrated any clinical or radiologic tumor response to vaccine therapy. Disease progression and death occurred in 10 of 11 patients at a median of 14 months (range, 3–23 months) and 19 months (range, 5–36 months) from diagnosis, respectively. All deaths were disease related. Patient 11 remained alive with stable disease 14 months after diagnosis.
To determine whether vaccination with DCs pulsed with tumor RNA was able to induce specific antitumor immunity, we used in vitro assays to examine both humoral and cell-mediated immune responses. We were unable to detect statistically significant cell-mediated antitumor responses in a T-cell proliferation assay (n = 5 patients) in any patients.
Enough tumor material was available to test humoral antitumor immunity by ELISA for the presence of tumor-specific antibodies in serum from three patients. Figure 2 shows that Patient 3 (P = 0.176) and Patient 4 (P = 0.015) had increases in specific antitumor antibodies after they received 3 administrations of DCRNA vaccine compared with pre-vaccine samples. Another patient (Patient 1) who was evaluated did not have measurable, specific, antitumor antibodies.
Quantitative lymphocyte cell counts are shown in Table 3. Total lymphocyte counts at the time of leukapheresis and in the PBMCs collected are shown in the first 2 columns of Table 3. On the day of the apheresis procedure, each patient (except Patient 1, who had slightly higher counts) had a total lymphocyte count that was within normal limits. One week prior to receiving the first DCRNA administration, total lymphocyte counts were within the normal range for all patients except Patients 3 and 11, who had below normal lymphocyte counts. Lymphocyte counts for this cohort of patients were higher than historic controls who did not receive lymphocyte-sparing chemotherapy (carboplatin). The average lymphocyte count for patients on this study (n = 6 patients) was 1390 × 106/L, whereas historic controls (n = 12 patients) at the same time point in treatment had an average lymphocyte count of 1075 × 106/L. This difference was not statistically significant due to the small sample numbers.
|Patient no.||Total lymphocyte count (× 106/L) (1200–3600)a||Lymphocyte phenotype (%)b|
|Day of apheresisc||PBMCs collectedd||Pre-DCRNA||CD3 (61–81)e||CD4 (32–56)e||CD8 (23–43)e||CD19 (7–19)e||CD56 (6–25)e|
Lymphocyte phenotyping data are available pre-DCRNA vaccine after other standard therapies were finished for all 7 vaccinated patients. CD3 cell counts generally were within normal limits, but T-lymphocyte subsets were abnormal for all patients. CD4 cell counts were below normal, and CD8 cell counts were above normal in all patients. Enumeration of CD19-expressing cells showed that all patients except Patient 9 had below normal counts. Two patients (Patients 2 and 4) had abnormal CD56 cell counts, whereas all other patients had normal CD56 counts. Overall, these data demonstrate abnormal lymphocyte subset distributions.
We examined lymphocyte proliferation in response to PHA at baseline and serially during DCRNA treatment. Immediately before the initiation of DCRNA therapy, all patients had significantly poorer proliferative responses to stimulation with PHA compared with normal controls (P = 0.003) (Fig. 3). However after receiving DCRNA vaccines, patients had slightly increased proliferation to PHA, although it continued at a significantly reduced level compared with normal controls (P = 0.001). Although patient responses to PHA after DCRNA vaccine generally were increased, the increase failed to achieve statistical significance (P = 0.07).
Figure 4 shows the lymphocyte proliferation in response to the superantigen staphylococcal enterotoxin B (SEB). Three of 4 patients who received ≥ 3 DCRNA vaccine administrations showed increased proliferation in response to SEB (P = 0.239).
Humoral immune parameters also were investigated pre-DCRNA vaccine. Table 4 shows titers of Ig isotypes at baseline, pre-DCRNA vaccine. IgG molecule numbers were within normal ranges for all patients tested. IgA and IgM isotypes were normal for all but 1 patient (Patient 11), who had below normal amounts for both Ig isotypes.
|Patient no.||Immunoglobulins (g/L)b||Tetanus (EU/mL) (0.16)c||Diphtheria (EU/mL) (0.10)c|
|IgG (5.18–17.8)||IgA (0.33–2.67)||IgM (0.32–1.35)||Pre||Post||Pre||Post|
|4||5.29||0.34||0.13||> 4.0||0.94||> 0.6||0.58|
Specific humoral responses to recall antigens, diphtheria, and tetanus are summarized in Table 4. Antibody titers were determined at enrolment and after immunization with DT booster vaccine and after receiving DCRNA. DT booster vaccine was given 1 week prior to receiving DCRNA vaccine. All patients had protective titers for tetanus prevaccine but generally were unable to boost titer levels after receiving vaccines except for Patient 9. It was shown that diphtheria titers prevaccine were at protective levels for 4 patients, whereas Patients 9 and 11 did not have protective titers. Patient 11 was unable to mount an increased response to diphtheria postvaccine.
The current study was designed to test the feasibility and safety of an autologous tumor RNA-pulsed, monocyte-derived DC antitumor vaccine in children with newly diagnosed neuroblastoma. We demonstrated that it is both safe and feasible to produce and administer this DCRNA vaccine preparation to this cohort of patients from a single apheresis. All patients tolerated the DCRNA vaccine well with no adverse reactions or clinical signs of toxicity or autoimmune responses reported. We were able to produce DCRNA vaccine preparations that fulfilled the feasibility requirements of 3 administrations each comprised of an IV and an ID component of 0.5 × 107 cells/m2 each for 8 of 9 apheresed patients.
We also examined the immune status of patients before they received the DCRNA vaccine to determine baseline immune function in this cohort. Cellular immune data from pre-DCRNA vaccine peripheral blood samples describe a cohort of patients with varying degrees of abnormality. Among these, the most striking shows that the normal ratio of CD4-positive and CD8-positive T cells was changed. It has been shown by others that CD4-positive T-cell regeneration lags behind CD8-positive T-cell regeneration after chemotherapy.15 The distribution of T cells in our patients showed that CD4-positive cell numbers were below normal for all patients, whereas CD8-positive cell numbers were above normal for all patients. In addition, CD19-positive cell numbers were below normal for all but 1 patient. Despite these irregularities, this cohort of patients generally showed good lymphocyte counts at their first vaccine administration approximately 6 weeks after completing intensive chemotherapy. Others have shown that lymphocyte counts remain below normal levels at 3 months after intensive chemotherapy.15 Although lymphocyte numbers improved in the current study, functional experiments for lymphocyte proliferation in response to PHA, as shown in Figure 3, demonstrated that cellular immune responses were impaired in these patients pre-DCRNA vaccine compared with the immune responses in normal controls.
Humoral responses demonstrate a similar pattern. Table 4 shows that, although most patients had normal Ig titers, only 3 patients were able to mount humoral responses to tetanus toxoid and diphtheria challenge. However, 2 patients were able to increase antitumor antibodies after the DCRNA vaccine, as shown in Figure 2. Again, these patients (Patients 3 and 4) received ≥ 3 vaccines, and it could be argued that their immune functions improved as the time after chemotherapy increased. In addition, Patients 3 and 4 had the longest progression-free survival, which may indicate that immune function was able to delay disease progression.
To our knowledge, this is the first study of its kind performed in patients with newly diagnosed neuroblastoma, and we showed here that this cohort of patients was immunocompromised profoundly prior to the intiation of DCRNA vaccine therapy. We recognized as part of this study the necessity of an intact and functional immune system for immunotherapy to be successful and therefore started patients on a “lymphocyte-sparing” regimen of chemotherapy prior to commencing traditional chemotherapy regimens. Although lymphocyte counts appeared to be improved over historic controls, immune function was compromised severely for both humoral and cellular responses prior to DCRNA vaccine. Given these results, it is not surprising that we were unable to detect robust antitumor responses in vitro.
Patient immunocompetency either before chemotherapy or after chemotherapy is clearly an area that is in need of further investigation to optimize immunotherapy for these patients. It has been demonstrated by others (for reviews, see Ingram and O'Rourke,29 Allan et al.,30 Swindle et al.,31 and Turtle et al.32) that cell-based immunotherapy using defined tumor antigens and defined HLA can elicit antitumor immune responses. Although this is an important advance in cancer therapy, it does not address the large number of tumors that lack defined tumor antigens. Therefore, approaches such as ours, using whole-tumor RNA as the source for pan tumor antigens, will require further investigation.
Our DCRNA vaccine preparation protocol was implemented prior to the realization that mature DCs generally are better stimulators of the antigen-specific immune response and therefore did not perform an additional maturation step after antigen uptake. To make a DC-based antitumor vaccine, it is necessary to produce immature DCs to take up antigen readily; once this occurs, DCs up-regulate costimulation molecule expression and down-regulate antigen uptake. Phenotypically mature DCs are defined as HLA-DRhigh, CD11c-positive, CD80-positive, CD86-positive, and CD1a-positive.33 Our DCRNA vaccine, however, did display some maturation molecule expression with phenotypes of HLA-DRhigh, CD86-positive, CD11c-positive, CD1a-negative, and a population of CD11c-negative cells (data not shown).
Progression-free and overall survival outcomes in this study were very poor, although they certainly fell within the range of experience for this disease. However, the number of patients on this study did not allow for any formal comparisons. Advanced stage neuroblastoma is a devastating disease that has a < 30% survival rate 3 years after diagnosis.2 The current Phase I trial demonstrated that, in a cohort of patients with newly diagnosed Stage 4 neuroblastoma, it was both safe and feasible to produce and administer monocyte-derived DCs that were pulsed with autologous tumor RNA. The immunocompetency of pediatric cancer patients is an area that has implications for both current clinical care and future treatments.
The authors acknowledge and thank Natalie Grapsas and Mary Brettell for their efforts in the clinical aspects of this trial. They also thank Dianne Tucker, Kerrie Clerici, and Michael Swain of the Cell Therapies Laboratory at The Royal Children's Hospital for the use of their facility and for helpful advice.