INTUVAX was manufactured at the Good Manufacturing Practice (GMP) compliant facility of the Cancer Centre Karolinska (CCK), Stockholm, Sweden. Monocytes were isolated from a leukocyte concentrate from a healthy blood donor at the Blood Centre, Karolinska University Hospital. The monocytes were cultivated in CellGro® DC Medium (CellGenix™), a serum free cell culture medium, supplemented with 100 ng/mL granulocyte macrophage colony-stimulating factor (GM-CSF) and 20 ng/mL interleukin 4 (IL-4) (both cytokines from CellGenix™). Thereafter, the cells were cultured in a closed system and incubated at 37 °C in 5% CO2 atmosphere. After differentiation, the CellGro® DC Medium was supplemented with 100 ng/mL GM-CSF and 20 ng/mL IL-4 and added to the cell suspension. To induce DC activation/maturation, toll-like receptor (TLR) 7/8 agonist R848 (2.5 μg/mL; InvivoGen), TLR3 agonist Poly I:C (20 μg/mL; Sigma-Aldrich), and human recombinant interferon gamma (IFN-γ; Imukin) (1000 U/mL; Boeringer-Ingelheim) were added to the culture medium. After maturation of DCs, mature pro-inflammatory DCs were harvested and resuspended in heat-inactivated AB plasma obtained from the Blood Centre, Karolinska University Hospital, and supplemented with 10% DMSO. One mL of the cell suspension was filled in each cryovials (CryoTubes™, NUNC) and subsequently deep-frozen to −150 °C, using a computerized gradient freezer (Planer) and thereafter transferred to a − 150 °C freezer. The final product, INTUVAX, was thawed and assessed for release tests (including testing for sterility, mycoplasma and endotoxin) for clinical use. All doses of INTUVAX that were used in this study originate from the same batch.
Men and women at least 18 years of age with newly diagnosed synchronous mRCC were enrolled. Inclusion required a size of the primary kidney tumor of at least 4 cm in longest diameter, as determined by CT, and at least one measurable distant metastatic lesion. Patients should have an Eastern Cooperative Oncology Group (ECOG) performance status <3 and were required to be candidates for nephrectomy and to have adequate hematological parameters. Therefore, patients with a life expectancy of less than 3 months, brain metastases, active or latent virus disease (HIV, HBV and HCV), other malignancy or ongoing active autoimmune disease, which required treatment with systemic immunosuppressive agents were excluded.
Study design and treatment
The study was a prospective single armed, open label phase I/II study. Patients on the waiting list for nephrectomy were enrolled in 3 cohorts of 3–5 patients per cohort. All patients in the same cohort were treated at the same dose level, i.e. 5 × 106, 10 × 106 or 20 × 106 viable and MHC class II expressing DCs. Two CT-guided injections with 14 ± 3 days of interval were administered into the viable part of the primary renal tumor identified with contrast enhanced CT scan. All INTUVAX doses used in the study were from the same batch.
The planned nephrectomy was performed within 28–42 days after the first vaccination. The patients were hospitalized for 24 h after each vaccination. The patients then returned to the clinic for safety visits one week after each vaccination, in connection with nephrectomy and 3 months after nephrectomy. Monitoring of vital signs, physical examination, safety lab and collection of adverse events (graded according to the National Cancer Institute Common Toxicity Criteria (CTC) version 4.0), were made at visits during the treatment period and at the follow-up visit, 3 months after nephrectomy.
Immunologic parameters including inflammatory immune serology, immune cell parameters, INTUVAX cell tracking and analyses using enzyme-linked immunosorbent spot (ELISpot) were monitored in connection with the vaccinations (samples taken at the day for vaccination and the day after). To evaluate potential auto- and allo-immune events, auto- and allo-immunization parameters were evaluated at screening and at follow-up visit 3 months after nephrectomy. Information on disease progression and survival data were obtained for all patients. Tumor follow-up, according to Response Evaluation Criteria In Solid Tumors (RECIST) 1.1 criteria, was conducted at baseline and at follow-up visit 3 months after nephrectomy. An extended follow-up to evaluate clinical outcome was further performed.
This study was approved by the local ethics committee in Uppsala, Sweden, and was conducted in accordance with Good Clinical Practice guidelines, as defined by the International Conference on Harmonisation. All patients provided written informed consent that was based on the principles of the Declaration of Helsinki. A safety monitoring committee reviewed safety during this study. The study was designed by the authors in collaboration with the sponsor (Immunicum AB, Gothenburg, Sweden).
The primary endpoints were registration of adverse events and changes in vital signs and laboratory parameters from baseline. Secondary end points included evaluation of tumor-specific immune response using immunohistology parameters of the renal tumor post nephrectomy, tumor-specific ELISpot assay, inflammatory immune parameters, systemic cytokine and chemokine production, immune cell distribution and activation, vaccine cell tracking and evaluation of potential auto- and allo-immunization. Finally, tumor response by CT scan was evaluated, according to RECIST 1.1.
Blood sample collection and storage
Heparinized blood was drawn before treatment, during the vaccinations, and at the end of the study. Peripheral blood mononuclear cells (PBMCs) were isolated by centrifugation on Metrizoate-ficoll (Lymphoprep, Nycomed/Axis-Shield PoC/Alere Technologies AS, Oslo, Norway) using a standard method, and thereafter aliquoted and cryopreserved for later use. Serum samples were also collected before treatment and at the end of the study and stored at −80 °C until analysis.
At screening, evaluation of the patient’s HLA-type (HLA-A, HLA-B and HLA-DR) was made. The samples were analyzed at the local clinical immunology laboratory at site (Dept of Clinical Immunology, Uppsala University Hospital, Uppsala, Sweden). HLA-typing of vaccine cell donor was also performed.
Tracking of injected INTUVAX cells
Flow cytometry analysis for donor cell tracking was performed on peripheral blood samples (taken just before vaccination and 1 h and 20–24 h after each vaccination) after lyzing of the erythrocytes with ammonium chloride and washing using a standard procedure. The samples were stained with combinations of murine monoclonal antibodies directly conjugated with fluorochromes (fluorescein isothiocyanate; FITC, phycoerythrin; PE, peridinin-chlorophyll protein; Per-CP or allophococyanin; APC). Staining was performed with antibodies specific for HLA class I antigens (HLA-A2 or HLA-A24) selectively expressed on INTUVAX cells, HLA-DR and CD86. The cells were incubated at 4 °C for 15 min with antibodies in the concentrations recommended by the manufacturer. The cells were analyzed in a FACSCanto™ II flow cytometer using the FACSDiva software and dot plots and quadrant statistics from a four-colour analysis were generated. All antibodies, the flow cytometer and the software were from BD Biosciences (Mountain View, CA, USA). Results for each subpopulation were expressed as the percentage of leucocytes. The samples were analyzed at the Dept of Clinical Immunology, Sahlgrenska University Hospital, Gothenburg, Sweden.
Serologic inflammatory parameters
Cytokines and chemokines were quantified from serum samples taken just before and 1 day after each administration of INTUVAX cells and 14 days after the second administration using a Bio-Plex human cytokine assay, a Bio-Plex 100 assay reader and the Bio-Plex Manager™ software (Bio-Rad Laboratories AB, Sundbyberg, Sweden). The analysis was performed according to the manufacturer’s instructions. The following inflammatory parameters were analyzed: IL-1 beta, IL-2, IL-4, IL-5, IL-6, IL-7, IL-8, IL-10, IL-12p70, IL-13, IL-17A, G-CSF. GM-CSF, IFN-gamma, MCP-1, MIP-1 beta and TNF-alpha. The chemokine RANTES was quantified by a commercially available ELISA (R&D Systems, Abingdon, UK), according to the manufacturer’s instructions. The production of TNF-alpha, IL-1 beta, IL-12p70, RANTES and MIP-1 beta was also analysed in supernatants from thawed and washed Intuvax cells cultured for 24 h in AIM-V culture medium at a concentration of 1 × 106 cells/mL. The samples were analyzed at the Dept of Clinical Immunology, Sahlgrenska University Hospital, Gothenburg, Sweden.
Lymphocyte distribution in peripheral blood
In order to evaluate potential changes in lymphocyte subset occurrence and activation state in peripheral blood taken just before and 1 day after each administration of INTUVAX cells and 14 days after the second administration, the following parameters were analyzed by FACS, as described above. Antibodies to the following antigens were used: CD3, CD4, CD8, CD19, CD16, CD56 and HLA-DR. The absolute number of blood lymphocytes was determined with Trucount reference beads using the method recommended by the manufacturer. The following subpopulations were reported CD3+, CD3 + 4+ and CD3 + 8+ T cells, CD3 + HLA-DR+ activated T cells, CD19+ B cells, CD3–16 + 56+ NK cells and CD3 + 16 + 56+ NKT cells. The results for each subpopulation were expressed as the percentage of lymphocytes and as the number of cells × 109/L. All antibodies and the Trucount beads were from BD Biosciences.
A randomly selected tissue specimen from the renal tumor was formalin-fixed, paraffin-embedded and cut into 5 μm sections. The sections were deparaffinised, and rehydrated in a graded series of ethanols. Antigen retrieval was done by heating tissue sections, using a Target Retrieval Solution, pH 9.0 (DAKO). The samples were subsequently incubated for 5 min in peroxidase blocking solution (DAKO) to inhibit endogenous peroxidase. Consecutive sections from each tumor specimen were then incubated with antibodies against CD3 (polyclonal rabbit anti-human CD3; DAKO), CD4 (monolonal mouse anti-human CD4, clone 4B12; DAKO), CD8 (polyclonal rabbit anti-human CD8; DAKO), CD56 (monoclonal mouse anti-human CD56, clone 123C3; DAKO), CD68 (monoclonal mouse anti-human CD68, clone KP1; DAKO), HLA-DR (monoclonal mouse anti-human HLA-DR, alpha chain, Clone TAL.1B5; DAKO) and PD-L1 (monoclonal mouse anti-human PD-L1, Clone 22C3, DAKO-kit SK006, DAKO) for 30 min. A subsequent reaction was carried out using secondary antibodies (DAKO) at 37 °C for 30 min. Then, the sections were washed three times with phosphate-buffered saline and subsequently the color was displayed with DAB (DAKO) for about 5 min. Sections were counterstained with haematoxylin, dehydrated in ethanol, cleared in xylene, and coverslipped.
For enumeration of CD8-infiltrating T cells, five different areas with most abundant positively stained cells in each section were selected and number of positve cells were counted under 400 magnification . The median number in each sample was then calculated. A semi-quantitative evaluation of infiltrating CD3+ T cells, CD4+ T cells, CD56+ NK/NKT cells, CD68+ macrophages and HLA-DR expressing non-tumor cells was performed. Moreover, the expression of HLA-DR by tumor cells was assessed.
The potential tumor-specific T cell response, induced by INTUVAX treatment was evaluated by running a tumor-specific IFN gamma (IFN-γ) ELISpot. Blood samples were collected at day 1 (before first vaccination) and 2 weeks after the second vaccination (in connection with hospitalization for nephrectomy). PBMC were isolated and frozen for subsequent batched analysis.
Fresh autologous tumor material from the resected tumor was deep-frozen (−70 °C) and subsequently thawed, homogenized with a GenltleMACS tissue homogenizer (Miltenyi Biotec GmbH, Bergisch Gladbach, Germany), followed by six cycles of freezing at −130 °C and thawing. After centrifugation and sterile filtration of the supernatant, the method of Bradford was used to determine protein concentration. The lysate was used as an antigenic source in the tumor-specific ELISpot assay.
Freshly drawn PBMCs from each patient were used for differentiation of immature monocyte-derived DCs, as described above for the production of INTUVAX. Immature DCs were subsequently pulsed with the tumor material (100 μg/mL) described above and stimulated for 24 h with a cocktail identical with the one used for INTUVAX production.
A direct ex vivo interferon gamma ELISpot analysis was then performed, using the commercial Human IFN-γ ELISpotPLUS kit (MABTECH AB, Nacka Strand, Sweden), according to instructions from the manufacturer.
Briefly, one vial of frozen PBMCs from each time point was thawed, counted in a cell counter (Sysmex K-4500; TOA Medical Electronics Co, Japan) and plated into the wells of an anti-IFN-γ antibody (clone 1-D1K, Mabtech, Inc) pre-coated ELISpot plate at 100,000 and 200,000 cells per well. Cells were then stimulated with 20,000 tumor-loaded autologous mature DCs per well for 24 h at 37 °C. As a positive control, an antibody to CD3 was used. After 24 h, plates were washed and incubated with a biotinylated anti-IFN-γ secondary antibody (clone 7-B6–1, Mabtech, Inc) for 2 h at room temperature. The plates were washed and incubated with streptavidin-conjugated alkaline phosphatase for 1 h and then washed again, and incubated with substrate for alkaline phosphatase. Excess substrate was removed by rinsing with tap water. The number of spots were counted using a dissection microscope. All samples were analyzed in triplicates, and the mean response (spots per 100,000 cells per well) was calculated. The analyses were performed at the Department of Clinical Immunology, Sahlgrenska University Hospital, Gothenburg, Sweden.
Measurement of potential auto- and alloimmunizaton
To evaluate potential autoimmune events, screening for autoantibodies against clinically relevant autoantigens, including nuclear antigens (ANA, SSA, SSB, Sm, RNP, Scl-70, Centromeres and Jo-1) and kidney parenchyma-associated autoantigens (liver-kidney microsomal antigens and mitochondrial antigens) was made. Samples were taken at screening and at day 120. The samples were analyzed at the Dept. of Clinical Immunology, Uppsala University Hospital, Uppsala, Sweden.
To evaluate potential vaccine-induced alloimmunization at the humoral level, screening for alloantibodies against HLA-A, B, C (MHC-class I) and HLA-DR, DQ, DP (MHC-class II) antigens was made. Samples were taken at screening and at day 120. The samples were analyzed at the Dept of Clinical Immunology at Uppsala University Hospital.
Tumor response was evaluated by CT scan, comparing the baseline CT scan (obtained within 28 days before day 1) with the CT scan made 3 months’ post nephrectomy, according to RECIST 1.1 criteria .