- EBV-B cells:
EBV-transformed B cells
Murine IgG2a Fc domain
During the last few years, HLA class I tetramers have been successfully used to demonstrate anti-vaccine CD8 CTL proliferation in cancer patients vaccinated with tumor antigens. Frequencies of CTL as low as 10–6 among CD8 cells were observed even in patients showing tumor regression. Little is known about the role of tumor-antigen-specific CD4 T cells in the context of these anti-vaccine responses. Therefore, we developed a very sensitive approach using fluorescent class-II–peptide multimers to detect antigen-specific CD4 T cells in vaccinated cancer patients. We produced HLA-DP4 multimers loaded with the MAGE-3243–258 peptide and used them to stain ex vivo PBL from melanoma patients injected with dendritic cells pulsed with several class I and class II tumor antigenic peptides, including the MAGE-3243–258 peptide. The multimer+ CD4 T cells were sorted and amplified in clonal conditions; specificity was assessed by their ability to secrete IFN-γ upon contact with the MAGE-3 antigen. We detected frequencies of about 1×10–6 anti-MAGE-3.DP4 cells among CD4 cells. A detailed analysis of one patient showed an anti-MAGE-3.DP4 CD4 T cell amplification of at least 3000-fold upon immunization. TCR analysis of the clones from this patient demonstrated a polyclonal response against the MAGE-3 peptide.
Among the available methods to characterize T cell immune responses, soluble MHC-class-I–peptide multimers have been widely used in clinical research to track and characterize antigen-specific CD8 CTL. In vaccinated cancer patients, a combination of in vitro re-stimulation of the PBL with the antigenic peptide and HLA class I multimer staining allowed the analysis, with high sensitivity and specificity, of anti-vaccine CTL responses at the clonal level1. These responses are usually present at low levels (∼1×10–4 to 1×10–7 of the CD8 cells) or sometimes undetectable even in patients who showed tumor regressions 1–4.
Although the importance of CTL in anti-tumor activity has been demonstrated, current protocols for active anti-cancer immunotherapy tend to include both HLA class I and class II-restricted antigens in the vaccine formulations with a view to enhance and sustain the anti-tumor responses 5, 6. Therefore, there is a need for reliable methods to enumerate and characterize the specificity and function of CD4 T cells induced in vaccinated cancer patients. Although the production of soluble MHC class I multimers has rapidly expanded since their first use in 1996 7, the development of MHC class II complexes has proved to be more difficult probably due to the intrinsic structural instability of soluble class II molecules. Recently however, some HLA-DQ and HLA-DR multimers have been obtained and evaluated for the detection of viral-antigen-specific CD4 T cells either ex vivo8–11 or after in vitro re-stimulation 10, 12–14. Recently, an approach for the ex vivo detection of CD4 T cells directed against a defined tumor antigen has been reported 15.
We have developed a method to analyze low-frequency tumor-antigen-specific CD4 T cell responses in vaccinated cancer patients by ex vivo staining of the PBL with HLA-class-II–peptide multimers, amplification of the sorted multimer+ CD4 clones and further assessment of their antigen specificity by functional assay. We have produced two different fluorescent HLA-DP4 multimers loaded with the antigenic peptide MAGE-3243–25816 and analyzed, with our ex vivo staining/cloning approach, the anti-MAGE-3.DP4 CD4 T cell responses in some patients vaccinated with autologous DC loaded with several HLA class I and class II tumor antigenic peptides, including the MAGE-3 peptide 5.
Production of soluble fluorescent MAGE-3.DP4–FcIg multimers
Soluble dimers of HLA-DP4 complexes loaded with the MAGE-3243–258 antigenic peptide (Fig. 1A) were produced in insect cells using the strategy previously described by Malherbe et al. 17. The MAGE-3 peptide, KKLLTQHFVQENYLEY, was covalently linked to the N terminus of the β chain DPB1*0401. The transmembrane regions of the α DPA1*0103 and β chains were replaced, respectively, by acidic and basic leucine-zipper peptides to facilitate pairing of the chains. The murine IgG2a Fc domain (FcIg) was added to the C terminus of the α chain to allow dimerization of the MAGE-3.DP4 complex and further purification of the dimer by affinity chromatography.
Fluorescent tetramers of the MAGE-3.DP4 complex were obtained by the addition of fluorescent protein A, which binds two FcIg portions. Usually, oligomers are produced because protein A has a tendency to form small aggregates. Although these oligomers were shown to stain cells of an anti-MAGE-3.DP4 CD4 clone (Fig. 2A), brighter staining was obtained when the MAGE-3.DP4–FcIg dimers were multimerized by the addition of a biotinylated anti-FcIg antibody and PE-labeled streptavidin (Fig. 2B).
The specificity of the MAGE-3.DP4–FcIg multimers was demonstrated by positive staining of the anti-MAGE-3.DP4 CD4 clone (Fig. 2B). As negative control, we labeled the cells of a CD4 T cell clone specific for the HLA-DR1-restricted MAGE-3267–282 peptide (Fig. 2C).
Optimal conditions for staining with the MAGE-3.DP4–FcIg multimers
Different parameters were evaluated to minimize the non-specific binding of the MAGE-3.DP4–FcIg multimers to cells from blood samples: (1) concentration of the multimers, (2) temperature and duration of incubation, (3) addition of antibodies specific of the T cell surface markers CD3, CD4 and CD8.
Staining of the anti-MAGE-3.DP4 CD4 T cell clone with the multimers indicated that a concentration of 50 nM resulted in a 50- to 100-fold increase of the mean fluorescence intensity compared with a non-relevant clone (Fig. 2B, C). This concentration was maintained in further experiments. Staining of the anti-MAGE-3.DP4 clone at different temperatures, demonstrated that the specific labeling was dramatically decreased at 4°C, whereas it was only slightly higher at 37°C than at room temperature (Fig. 2D). Incubation at room temperature for 1 h was found to be adequate to stain our anti-MAGE-3.DP4 CD4 clone (Fig. 2E). However, we do not exclude that other anti-MAGE-3.DP4 CD4 clones with low TCR affinity would require higher temperature and/or prolonged incubation time for positive staining 18.
We next investigated whether the addition of anti-CD4 antibodies could interfere with the binding of the multimers to the anti-MAGE-3.DP4 CD4 clone. Two different monoclonal antibodies – Multi-clone and SK3 – were tested and were shown to reduce significantly the multimer staining (Fig. 2F). On the contrary, anti-CD3 antibody, UCHT1, marginally affected the binding of the multimers (data not shown). On the basis of these findings, anti-CD3 UCHT1 and anti-CD8 were used in the following experiments to label the PBL and select the CD3+CD8–multimer+ cells.
Non-specific binding of the MAGE-3.DP4–FcIg multimers to PBL was then evaluated on lymphocytes from a non-cancerous DP4+ patient. Cells were first labeled with 50 nM of the MAGE-3.DP4–FcIg multimers and then with anti-CD3 and anti-CD8 antibodies. The temperature and duration of incubation were varied. The results indicated that incubation at room temperature for 1 h gave a low background (0.05% of CD3+CD8– cells), which is twice as high after 2 h of incubation. Incubation at 37°C for 1 h significantly increased the non-specific binding of the multimers (0.23%). The lowest background was observed at 4°C (0.01%). Considering the significant mean fluorescence intensity decrease of the anti-MAGE-3.DP4 clone stained with the multimers at 4°C (Fig. 2D) and the high background generated by staining of the PBL at 37°C, we defined the following conditions for PBL labeling: incubation for 1 h at room temperature with 50 nM of the multimers, followed by staining for 10 min at room temperature with the anti-CD3 and anti-CD8 antibodies.
Evaluation of the anti-MAGE-3.DP4 CD4 T cell response in vaccinated cancer patients
We analyzed the anti-MAGE-3.DP4 CD4 T cell response in some melanoma patients vaccinated with DC pulsed with several HLA class I and class II peptides, including the MAGE-3243–258 peptide 5. Among the 16 fully evaluable patients, one month after the fifth injection, 7 showed disease progression, 8 exhibited stable disease and 1 experienced complete regression 5.
The approach we developed to monitor the anti-MAGE-3.DP4 CD4 T cell response is outlined in Fig. 3. Briefly, frozen PBMC were thawed and kept overnight in culture medium. Cells were collected and labeled with MAGE-3.DP4–FcIg multimers, anti-CD3 and anti-CD8 antibodies. Multimer+CD3+CD8– cells were isolated by FACS and stimulated by DP4+ EBV-transformed B cells (EBV-B cells) pulsed with the MAGE-3 peptide. After two weekly re-stimulations, the specificity of the proliferating clones was defined by their ability to produce IFN-γ upon contact with EBV-B cells exogenously loaded either with the MAGE-3.DP4 peptide or a bacterial recombinant MAGE-3 protein. The clones were also tested for recognition of EBV-B cells transduced with a retrovirus coding for the MAGE-3 protein fused to a truncated invariant chain (Ii–MAGE-3) to target the protein to the endosomal compartment. Finally, the IFN-γ-secreting clones were tested for staining with the MAGE-3.DP4–FcIg multimers. The frequency of anti-MAGE-3.DP4 CD4 T cell clones among CD4 blood T lymphocytes was estimated with the formula annotated in Fig. 3.
PBL from patient R12 5 were analyzed before immunization and after the second, fifth and ninth immunization (post-2, -5, -9). Patient R12 showed stable disease one month after the fifth injection. No anti-MAGE-3.DP4 CD4 T cell could be detected in pre-immunization PBL leading to a frequency estimate below 2.6×10–7 of CD4 blood T lymphocytes. Several clones, secreting IFN-γ specifically upon contact with the antigen (Fig. 4A), were obtained in post-vaccination samples. All of them stained with the MAGE-3.DP4–FcIg multimer (Fig. 4B). The level of the anti-MAGE-3.DP4 response significantly increased after the first two injections and peaked after the fifth injection, to a frequency of 7.5×10–4 (Fig. 5A). This represented at least a 3000-fold increase of the frequency found before immunization.
We analyzed the post-5 blood sample of another patient, R15, who also showed stable disease one month after the fifth injection. We observed a frequency of anti-MAGE-3.DP4 CD4 T cells of about 3×10–6.
Finally, we analyzed pre- and post-5 blood samples of patient R02. This patient was injected with DC pulsed with the same peptide mixture as patient R12, but the MAGE-3.DP4 peptide was omitted. Anti-MAGE-3.DP4 CD4 T cells were detected at a frequency of 1.2×10–6 and 1.5×10–6 in pre- and post-5 PBL respectively, indicating that MAGE-3.DP4 T cell precursors were present before DC immunization and were not induced to proliferate after vaccination.
Functional characterization of the MAGE-3.DP4 CD4 clones from patient R12
We first evaluated the ability of the anti-MAGE-3.DP4 clones isolated from patient R12 to recognize DP4+ tumor cells expressing MAGE-3. We demonstrated previously that the MAGE-3.DP4 peptide could be naturally presented at the surface of tumor cells 16. Ten different clones were tested. All of them were stimulated to secrete IFN-γ upon contact with two melanoma cell lines, as illustrated in Fig. 4C for two clones. The cytokine profile of these two clones was typical of a Th1 type phenotype as they produced high levels of IFN-γ and TNF-α and almost no IL-4 and IL-10, when stimulated with DP4+ EBV-B cells loaded with the MAGE-3 peptide (Fig. 4D). Although we cannot exclude that the conditions we used to amplify the clones have introduced a bias towards this phenotype, this result is in accordance with the strong Th1 response detected by ELISPOT in the PBL of this patient 5.
The anti-MAGE-3.DP4 CD4 T cell immune response of patient R12 is polyclonal
To evaluate the diversity of the anti-MAGE-3.DP4 CD4 immune response in patient R12, the TCR-β usage of 21 randomly selected clones was determined. Sixteen different TCRVβJβ clonotypes (Fig. 5B) were found with around 50% of the clones sharing the same Vβ gene, Vβ12. This clearly revealed a CD4 polyclonal immune response against the MAGE-3.DP4 peptide.
Improved detection of anti-MAGE-3.DP4 CD4 T cells by using multimers generated from biotinylated MAGE-3.DP4 monomers
Although the above-described procedure allowed us to detect anti-MAGE-3.DP4 CD4 T cells with frequencies as low as 1.2×10–6 of the CD4 cells, we investigated the possibility to further reduce the non-specific binding of the multimers to PBL stained ex vivo. We suspected that part of the background was inherent to the use of the biotinylated anti-FcIg antibody for the multimerization of the MAGE-3.DP4–FcIg dimers. We thus set up the construction of the multimers from biotinylated MAGE-3.DP4 monomers. The structure of the MAGE-3.DP4 monomers was similar to the MAGE-3.DP4–FcIg construct with the following differences (Fig. 1B). The murine FcIg portion of the recombinant α chain was exchanged for a sequence of six histidines and a tag for biotinylation; a sequence of six histidines was also fused to the C terminus of the recombinant β chain.
Monomers were purified by immobilized metal ion chromatography and biotinylated in vitro. Fluorescent multimers were produced by the addition of PE-labeled streptavidin. Specificity of the MAGE-3.DP4 multimers was confirmed by their ability to stain the anti-MAGE-3.DP4 clone but not the irrelevant anti-MAGE-3.DR1 clone (Fig. 6). By testing additional anti-CD4 antibodies, we identified clone RPA-T4, which did not affect binding of the multimer (Fig. 6). Staining of PBL from a non-cancerous patient with the MAGE-3.DP4 multimers showed a 5-fold lower level of non-specific binding (0.01% of the CD4 cells) in comparison to the MAGE-3.DP4–FcIg multimers (0.05%) (data not shown).
We compared the two different MAGE-3.DP4 multimers by analyzing the anti-MAGE-3.DP4 CD4 T cell response in a post-2 blood sample from patient R12. PBL from the same sample were labeled ex vivo with either the MAGE-3.DP4–FcIg multimer (+ anti-CD3, anti-CD8) or the MAGE-3.DP4 multimer (+ anti-CD4, anti-CD8). Multimer+ CD4 cells were cloned, amplified by in vitro re-stimulation and tested for specificity both by IFN-γ assay and multimer staining. The results are summarized in Table 1. Frequency estimates of anti-MAGE-3.DP4 CD4 T cells were similar for the two multimers but the efficiency of the detection was significantly increased (∼7-fold) by staining with the MAGE-3.DP4 multimers.
|Multimers||Cells introduced in FACS||% of multimer+ CD4 cells||Sorted cells||Cloned cells||Proliferating clones||Anti-MAGE-3.DP4 clones||Efficiency of detection (%)d)||Frequency of anti-MAGE-3.DP4 clones|
Here we describe the production of HLA-DP4 multimers loaded with the tumor-specific antigenic peptide MAGE-3243–258 and a strategy to monitor the anti-MAGE-3.DP4 CD4 T cell responses in PBL of some vaccinated melanoma patients.
In order to analyze these CD4 T cell responses, we felt it crucial to characterize at the clonal level the CD4 T cells labeled with the multimers. By combining ex vivo staining of the PBL and cloning of the multimer+ cells, we were able to detect about one anti-MAGE-3.DP4 CD4 T cell among 106 CD4 cells, starting with frozen lymphocyte samples. The background of multimer+ cells was minimized by optimizing the conditions for staining and by using multimers generated from biotinylated MAGE-3.DP4 monomers. In addition, strict specificity of the analysis was achieved through the examination of the reactivity of the clones against the naturally processed MAGE-3 antigen.
The frequency of the anti-MAGE-3.DP4 CD4 T cell response was examined over time for patient R12 and compared with the results obtained previously by IFN-γ ELISPOT assay (Table 2 and unpublished results) 5. Although the levels of the post-vaccine responses determined by both methods were similar, the estimate of the pre-immunization frequency obtained by multimer detection was more than 100-fold lower than that evaluated by ELISPOT (Table 2).
|Patient||Blood sample||Frequency among CD4 T cells|
This difference may reflect the limits in sensitivity of the ELISPOT assay which is reported to be in the range of 1×10–4 of the CD4 lymphocytes, but we cannot exclude formally that this discrepancy results from the well-known observation that some class II peptides can bind to different HLA molecules. This is illustrated by the analysis of patient R15. The frequency of anti-MAGE-3 CD4 T cells determined in post-5 PBL by the DP4 multimer analysis was 10 times lower than by ELISPOT (Table 2). However, Schultz et al. 19 showed that the CD4 T cell clones they isolated from this patient presented the MAGE-3 peptide in association with the HLA molecule DQ6, but not DP4. Taken together, our results demonstrated that the multimer staining/cloning procedure displayed higher sensitivity than the ELISPOT assay but could underestimate the CD4 T cell responses against the vaccine peptide when it binds to HLA class II molecules that were not identified previously.
One advantage of the multimer staining approach is that antigen-specific CD4 T cells are detected independently of their effector functions. In our analysis, several randomly selected anti-MAGE-3.DP4 clones isolated from patient R12 were shown to secrete IFN-γ after contact with DP4 tumor cell lines expressing the MAGE-3 antigen, suggesting that these clones could be involved in the anti-tumor response by direct recognition of the melanoma cells at the tumor site. Our procedure also allows the detection of anti-vaccine CD4 T cells secreting other cytokines than IFN-γ, including for instance CD4+CD25+ T regulatory cells, which might suppress T cell responses against the tumor 20.
The TCR analysis of the anti-MAGE-3.DP4 CD4 T cell clones showed that patient R12 developed a polyclonal response. Polyclonality of anti-vaccine CD8 T cell responses has also been observed in patients immunized with DC pulsed with the HLA class I peptide MAGE-3.A1 21, in contrast to the monoclonal responses induced in patients vaccinated with the MAGE-3.A1 peptide or with a recombinant ALVAC virus expressing the MAGE-3.A1 and MAGE-1.A1 epitopes 1, 4, 22. Thus, our results support the idea that vaccination with DC results in the activation of more T cell precursors.
In conclusion, we show here that our combined ex vivo multimer staining/cloning approach is a reliable method to follow at the clonal level low-frequency antigen-specific CD4 T cell responses in cancer patients. We present here a first detailed analysis of the frequency and TCR diversity of the anti-MAGE-3.DP4 CD4 T cell response in a vaccinated cancer patient. Further analyses have to be performed to define the role of these anti-vaccine CD4 T cells in anti-tumor responses.
Materials and methods
Melanoma patients R2, R12 and R15
Melanoma patients R2, R12 and R15 were included in a clinical trial in which mature monocyte-derived DC loaded with multi-HLA class I and class II peptides were administered subcutaneously at 14-day intervals for the first five injections and then at 1-month intervals 5. In the case of HLA class II peptides, DC were loaded with MAGE-3243–258.DP4, MAGE-3121–134.DR13, tyrosinase.DR4, or gp100.DR4 peptides. Blood cells were collected before immunization and at different time points after immunization. PBMC were then isolated and cryoconserved.
Cell lines, media, and reagents
The EBV-B cells and the melanoma cell lines MZ2-MEL.43 and MI13443 were cultured in IMDM (Gibco BRL, Gaithersburg, MD, USA) supplemented with 10% FCS (Gibco BRL), 0.24 mM L-asparagine, 0.55 mM L-arginine, 1.5 mM L-glutamine (AAG), 100 U/ml penicillin and 100 μg/ml streptomycin. Human recombinant IL-2 was purchased from Eurocetus (Amsterdam, The Netherlands), IL-7 from Genzyme (Cambridge, MA, USA). Human recombinant IL-4, IL-6, and IL-12 were produced in our laboratory. The MAGE-3 recombinant protein was produced in Escherichia coli as described by Zhang et al.23 and provided by GlaxoSmithKline Biologicals (Rixensart, Belgium). Antibodies anti-CD4–FITC (Multi-Clone LeuTM-3a+3b, SK3 and RPA-T4), anti-CD8–FITC, anti-CD8–Cy5PE and anti-CD3–allophycocyanin (APC) (UCHT1) were purchased from BD Biosciences (Mountain View, CA, USA).
IFN-γ ELISPOT assay
PBMC were added in triplicates at 5×105 cells per 96-well precoated with anti-IF-γ mAb (1-D1K; Mabtech, Hamburg, Germany). Then KLH or MAGE-3.DP4 peptide were added at 10 μg/ml and, after 20 h, wells were washed and incubated with biotinylated mAb to IFN-γ (7-B6-1; Mabtech) for 2 h. Final staining and computer-assisted analysis was done as described previously 24, 25. Background without antigen was less than three spots. Responses were considered significant if a minimum of five spot-forming cells per well were detected and, additionally, this number was at least twice that in negative control wells.
Transduction of EBV-B cells with the retrovirus encoding Ii–MAGE-3
The retroviral vector for expression of the Ii–MAGE-3 gene was constructed and transfected into EBV-B cell lines as described previously 23.
Production of soluble fluorescent MAGE-3.DP4–FcIg multimers
Soluble MAGE-3.DP4–FcIg dimers were produced in Drosophila S2 cells essentially as described by Malherbe et al. 17. HLA-DPA1*0103 α and DPB1*0401 β chains were truncated to remove their transmembrane and cytosolic regions. Acidic and basic leucine-zipper peptides were added, through a five-amino-acid linker (GGGGS), at the C terminus of the α and β chain, respectively, to facilitate pairing of the chains 26. The MAGE-3.DP4 peptide KKLLTQHFVQENYLEY was covalently introduced, with a flexible 16-amino-acid linker (GGGGSLVPRGSGGGGS) at its C terminus, between the third and the fourth N-terminal amino acid of the β chain. The α chain was further modified by the addition of the murine IgG2a Fc domain to allow dimerization of the MAGE-3.DP4 complex 27.
The recombinant genes were cloned in the pRMHA3 expression vector 28 and cotransfected in Drosophila S2 cells with a neomycin-resistance gene to produce stable transfectants. Soluble MAGE-3.DP4–FcIg complexes were purified from the supernatant of Drosophila S2 transfected cells by protein G affinity chromatography and subsequent ion-exchange chromatography. Fluorescent complexes were obtained either by the addition of an Alexa Fluor®488-conjugated protein A (Molecular Probes, Leiden, The Netherlands) or biotin-conjugated rat anti-mouse IgG2a Fc (BD Biosciences Pharmingen, Erembodegem, Belgium) and R-PE-conjugated streptavidin (BD Biosciences Pharmingen). The concentration of the multimer used for cell staining is expressed as the concentration of the MAGE-3.DP4 monomer in the sample.
Production of soluble fluorescent MAGE-3.DP4 multimers
Soluble biotinylated MAGE-3.DP4 complexes were produced as described for the MAGE-3.DP4–FcIg dimers with the following differences. The α chain was modified by exchange of the murine IgG2a Fc domain for a six-histidine tag to allow further purification of the complex. A biotinylation tag followed by a six-histidine tag was introduced at the C terminus of the basic leucine-zipper peptide of the recombinant MAGE-3.DP4 β chain. The recombinant genes were cloned in the pRMHA3 expression vector and cotransfected in Drosophila S2 cells. Soluble monomers were purified from the supernatant of Drosophila S2 transfected cells by immobilized metal ion chromatography and biotinylated in vitro with the birA enzyme (Avidity, Denver, CO, USA). The biotinylated monomers were further purified by gel-filtration chromatography. Fluorescent multimers were produced by the addition of PE-labeled streptavidin. The concentration of the multimer used for cell staining is expressed as the concentration of the MAGE-3.DP4 monomer in the sample.
T cell reference clones
The anti-MAGE-3.DP4-specific T cell clone, R12/C9, was used as a positive reference clone. This clone was obtained in our laboratory using the IFN-γ Secretion Assay (Miltenyi Biotech, Bergisch Gladbach, Germany). Briefly, PBMC from melanoma patient R12 were stimulated with 10 μg/ml of the MAGE-3.DP4 peptide KKLLTQHFVQENYLEY for 16 h. Then the cells were labeled with anti-CD4 and anti-IFN-γ antibodies. IFN-γ-secreting CD4+ T cells were sorted by a FACSVantage™ flow cytometer (BD Biosciences).
The sorted cells were cloned and stimulated by the addition (1×104 cells/well) of irradiated allogeneic HLA-DP4 EBV-B cells (obtained from melanoma patient DDHK2 23) pulsed with 5 μg/ml of the MAGE-3.DP4 peptide in the presence of cytokines IL-2 (50 U/ml), IL-4 (5 U/ml), IL-7 (10 ng/ml) and phytohemagglutinin (PHA) (125 ng/ml). Another irradiated allogeneic EBV-B cell line, LG2-EBV, was added as a source of feeder cells (1×104 cells/well). After 28 days, each growing clone (∼5×103 cells) was tested for specificity by stimulation with ∼2×104 DDHK2 EBV-B cells either pulsed with 5 μg/ml of MAGE-3.DP4 peptide or expressing retro-Ii–MAGE-3. Clone R12/C9 specifically recognized the MAGE-3 peptide restricted by HLA-DP4. The DDHK2 clone 1, used as negative control clone, was derived from melanoma patient DDHK2 and recognized the MAGE-3 peptide ACYEFLWGPRALVETS restricted by HLA-DR1 23.
Staining of PBL for ex vivo sorting with the MAGE-3.DP4–FcIg and MAGE-3.DP4 multimers
PBMC from vaccinated patients were thawed, and incubated at 37°C overnight at 4×106 cells/ml in Iscove's medium containing 10% AB human serum (HS) in the presence of DNAse (5 U/ml). The next day, cells were collected, washed, resuspended at 107/ml in PBS with 1 mM EDTA and 1% HS (PBS-HS 1%) and first incubated for 1 h at room temperature with 50 nM of the R-PE-labeled MAGE-3.DP4–FcIg or MAGE-3.DP4 multimers (concentrations refer to the concentration of the MAGE-3.DP4 monomers in the sample). For the MAGE-3.DP4–FcIg multimer staining, anti-CD3–APC (UCHT1; 1/40) and anti-CD8–FITC, (1/40) antibodies were subsequently added to the cells and incubation was continued for 10 min. For the MAGE-3.DP4 multimer, cells were stained in the same conditions with anti-CD4–FITC RPA-T4 (1/40) and anti-CD8–Cy5PE (1/40). The cells were washed, resuspended in PBS-HS 1% and sorted by the FACSVantage flow cytometer. Cells were gated on living CD3+CD8–multimer+ or living CD4+CD8–multimer+ cells, respectively, and cloned at one cell/well.
Culture of ex vivo cloned cells
Cloned cells were weekly stimulated by the addition of irradiated DDHK2 EBV-B cells (1×104 cells/well) pulsed with the MAGE-3.DP4 peptide (5 μg/ml). LG2-EBV was added as a source of feeder cells (1×104 cells/well), in Iscove's medium containing 10% AB HS and AAG, with the addition of IL-2 (50 U/ml), IL-7 (10 ng/ml), and PHA (125 ng/ml).
MAGE-3.DP4 specificity assay of the clones
After two weekly re-stimulations, the cells were kept resting for two weeks. In total, 5,000 cells of each clone were stimulated with 2×104 DDHK2 EBV-B cells either pulsed for 4 h with 5 μg/ml of the MAGE-3.DP4 peptide or loaded for 20 h with 20 μg/ml of recombinant protein MAGE-3, or expressing retro-Ii–MAGE-3. After 20 h of co-culture in round-bottomed 96-well plates with 150 μl/well complete IMDM supplemented with IL-2 (25 U/ml), IFN-γ released in the supernatant was measured by ELISA using reagents from Medgenix Diagnostics-Biosource (Fleurus, Belgium). The release of IFN-γ had to be at least two times and exceed by more than 150 pg/ml the release of IFN-γ upon contact with control DDHK2 EBV-B cells.
Recognition of tumor cells
Tumor cells were distributed at 20,000 cells per round-bottomed 96-well with 5,000 CD4+ T lymphocytes in 150 μl of complete IMDM supplemented with IL-2 (25 U/ml). Supernatants were harvested after 20 h of co-culture and IFN-γ production was measured by ELISA.
Analysis of the cytokine profile of the clones
DDHK2 EBV-B cells (20,000) were pulsed with 5 μg/ml of the MAGE-3.DP4 peptide and then co-cultured overnight with 5,000 CD4+ T cells in a total volume of 150 μl of IMDM supplemented with 10% HS. Supernatants were harvested and stored at –80°C until cytokine testing was performed. IL-2, IL-4, IL-5, IL-10, TNF-α, and IFN-γ were detected by the Cytometric Bead Array kit (BD Biosciences Pharmingen, San Diego, CA, USA).
RNA was extracted from 3×105 cells of each MAGE-3.DP4 CD4 T cell clone with the Tripure reagent (Boehringer Mannheim, Mannheim, Germany), and subsequently converted to cDNA at 42°C for 90 min with 200 U M-murine leukemia virus reverse transcriptase (Life Technologies, Merelbeke, Belgium). The TCR Vβ usage was assessed by PCR amplification by using a complete panel of Vβ-specific sense primers and Cβ anti-sense primer, respectively 29. Primers were chosen on the basis of described panels of TCR V-region oligonucleotides, and with alignments of TCR sequences available at the International Immunogenetics Database (http://imgt.cines.fr). Each PCR product was purified and sequenced to obtain a full identification of the CDR3 region.
We thank Mrs S. Ottaviani and C. Wildmann for their precious assistance, and Mrs N. Krack for editorial assistance. Yi Zhang was supported by a postdoctoral fellowship from the International Institute of Cellular Pathology, Brussels, Belgium. Nikolina Renkvist was supported by a post-doctoral fellowship from the Training and Mobility of Researchers program of the European Commission. This work was supported by the Belgian Programme on Interuniversity Poles of Attraction initiated by the Belgian State, Prime Minister's Office, Science Policy Programming, and by a grant from the Fédération Belge contre le Cancer (Belgium). The clinical trial was supported by a “Cancer Research Institute Melanoma Initiative Clinial Trial Grant” to Gerold Schuler.