Efficient ex vivo generation of dendritic cells from CD14+ blood monocytes in the presence of human serum albumin for use in clinical vaccine trials

Authors


Naoyuki Katayama MD, The Second Department of Internal Medicine, Mie University School of Medicine. 2–174 Edobashi, Tsu, Mie 514–8507, Japan. E-mail: n-kata@clin.medic.mie-u.ac.jp

Abstract

Dendritic cells (DC) with the potential to induce anti-tumour immunity represent one of the promising candidates for cancer vaccines. Efficiency of ex vivo DC generation depends on culture conditions, especially protein components in the plasma or serum used. Using human serum albumin (HSA), we devised a constant and reproducible culture method for DC generation from peripheral blood CD14+ cells. The number of DC obtained with 2% HSA-supplemented cultures containing granulocyte-macrophage colony-stimulating factor and interleukin 4 were consistently higher than in cultures with various concentrations of autologous plasma or serum. The concentrations and time points tested for plasma or serum considerably affected the number of DC recovered. DC prepared with HSA acquired the ability to uptake dextran, and expressed high levels of major histocompatibility (MHC) and co-stimulatory molecules similar to DC cultured with autologous plasma or serum. Although DC cultured with autologous plasma or serum consisted of CD1a+ and CD1a populations, DC differentiated in the presence of HSA expressed CD1a. DC obtained with HSA primed and induced immunogenic peptide-specific cytotoxic T lymphocytes against a tumour rejection antigen, HER2. These findings suggest that our method for preparation of DC with HSA should prove valuable in DC generation for immunotherapy.

Understanding the molecular events involved in immune recognition by T cells has provided opportunities to treat cancer patients with vaccines that induce potent antigen-specific T-cell responses (Pardoll, 1998). Recent studies have identified a number of tumour antigens, most of which are targeted by cytotoxic T lymphocytes (CTL) (Houghton, 1994; Boon et al, 1995; Van den Eynde & van der Bruggen, 1997). Several tumour antigen-derived peptides, which are presented by common major histocompatibility (MHC) class I molecules on antigen-presenting cells (APC) and proved to be immunogenic, have also been defined (Van den Eynde & van der Bruggen, 1997). As the simple administration of peptide would lead to tolerance rather than to immune rejection (Aichele et al, 1995; Toes et al, 1996), the use of professional APC, either loaded with immunogenic peptides on their empty MHC molecules or pulsed with tumour lysate, may represent promising candidates for therapeutic cancer vaccines (Toes et al, 1998).

Dendritic cells (DC) are APC distinguished by their ability to initiate primary T cell-mediated immune responses (Steinman, 1991; Caux et al, 1995; Cella et al, 1997; Hart, 1997; Banchereau & Steinman, 1998; Banchereau et al, 2000). DC display dramatic changes in functions during their maturation. Immature DC capture and process antigens, whereas mature DC prime and activate naive T cells (Steinman, 1991; Caux et al, 1995; Cella et al, 1997; Hart, 1997; Banchereau & Steinman, 1998; Banchereau et al, 2000). In clinical cell-based cancer vaccine approaches, DC have been used as a source of adjuvants required to induce immunity against tumour-specific or tumour-selective antigens (Hsu et al, 1996; Murphy et al, 1996; Nestle et al, 1998; Lim & Bailey-Wood, 1999; Morse et al, 1999; Reichardt et al, 1999; Thurner et al, 1999; Mackensen et al, 2000; Titzer et al, 2000). One major focus has been on efficient ex vivo generation of DC with the potential to induce immune responses, especially elicited by specific CTL, against tumour rejection antigens expressed by the tumour. Practically, autologous DC have been either isolated directly from leukapheresis products (Hsu et al, 1996; Reichardt et al, 1999) or generated in culture from adherent peripheral blood mononuclear cells (PBMCs) (Murphy et al, 1996; Nestle et al, 1998; Lim & Bailey-Wood, 1999; Morse et al, 1999), blood monocytes (Thurner et al, 1999) or mobilized CD34+ peripheral blood haemopoietic progenitor cells (Mackensen et al, 2000; Titzer et al, 2000) in clinical DC-based vaccine trials. In direct isolation from leukapheresis products, the yield and purity of DC prepared vary considerably among patients (Hsu et al, 1996; Reichardt et al, 1999). DC generation from mobilized CD34+ peripheral blood haemopoietic progenitor cells relies on the treatment of patients with haemopoietic growth factors such as granulocyte colony-stimulating factor (G-CSF) as well as the leukapheresis (Mackensen et al, 2000; Titzer et al, 2000). The culture of adherent PBMCs or blood monocytes is a simple method but commonly needs the support of plasma or serum (Bender et al, 1996; Murphy et al, 1996; Romani et al, 1996; Jonuleit et al, 1997; Nestle et al, 1998; Lim & Bailey-Wood, 1999; Thurner et al, 1999; Duperrier et al, 2000). Because serum or plasma contains growth factors or other substances, both of which may influence the generation of DC (Siena et al, 1995; Mitani et al, 2000), the numbers of DC obtained from cultures supplemented with plasma or serum would not be predictable. Therefore, it is necessary to test whether appropriate numbers of DC can be obtained in the individual patients during screening of eligibility for DC-based vaccine therapy. Serum-free media have been used for generation of DC from apheresis cells (Tarte et al, 2000), haematopoietic progenitors (Mitterer et al, 1998), PBMCs (Chen et al, 1998) and monocytes (Duperrier et al, 2000). However, the serum-free media used in those studies contained human-derived proteins, which may modify the function of DC.

In the present study, we developed a fully defined serum-free culture system for the generation of DC from human CD14+ blood monocytes. Our defined culture condition contains granulocyte-macrophage colony-stimulating factor (GM-CSF) and interleukin 4 (IL-4) as cytokines and human serum albumin (HSA) as the protein components. Our culture procedure, described here, should be useful for stable generation of DC in clinical trials.

Materials and methods

Media and reagents The medium used was Roswell Park Memorial Institute (RPMI)-1640 medium (Nissui Pharmaceutical, Tokyo, Japan) supplemented with 2 mmol/l l-glutamine, 50 U/ml penicillin, 50 µg/ml streptomycin and either 10% fetal calf serum (FCS) (Hyclone Laboratories, Logan, USA), 0·1–10% human plasma or serum or 0·1–10% HSA (Albumin Cutter; Bayer Pharmaceutical, Osaka, Japan). Ten batches of HSA from Bayer Pharmaceutical, five batches of HSA (Buminate) from Baxter Healthcare (Tokyo, Japan) and five batches of HSA (Albumin-Nichiyaku) from Nihon Pharmaceitical (Tokyo, Japan) were screened to test their ability to support DC generation, as described below. Purified recombinant human GM-CSF was kindly provided by Kirin Brewery, Tokyo, Japan. Purified recombinant human IL-4 was a generous gift from Ono Pharmaceutical, Osaka, Japan. Purified recombinant human tumour necrosis factor-α (TNF-α) was provided by Dainippon Pharmaceutical, Suita, Japan and purified recombinant human IL-2 was a gift from Takeda Pharmaceutical, Osaka, Japan. Concentrations of growth factors used in this study were as follows: GM-CSF, 10 ng/ml; IL-4, 10 ng/ml; TNF-α, 20 ng/ml; IL-2, 20 IU/ml.

Antibodies In this study, mouse monoclonal antibodies (mAbs) to the following human molecules were used: fluorescence isothiocyanate (FITC)-conjugated mAb specific for CD1a (Ortho Diagnostic System, Raritan, USA); phycoerythrin (PE)-conjugated mAbs specific for CD1a and CD40 (Immunotech, Marseille, France); FITC- or PE-conjugated mAbs specific for CD14, CD54, CD80 and HLA-DR (Becton Dickinson, San Jose, USA); PE-conjugated mAb specific for HLA-A,B,C (DAKO, Glostrup, Denmark); PE-conjugated mAb specific for CD86 (PharMingen, San Diego, USA.); FITC-conjugated mAb specific for myeloperoxidase (MPO) (DAKO). FITC- or PE-conjugated mouse IgG1, IgG2a (Becton Dickinson) or IgG2b (Coulter, Hialeah, USA) served as an isotype control.

Cell preparation After receiving informed consent, peripheral blood was obtained from healthy adult Japanese volunteers in heparinized syringes. PBMCs were separated using Ficoll–Hypaque (Nycomed Pharma AS, Oslo, Norway) density gradient centrifugation. CD14+ cells were isolated from PBMCs using the MACS CD14 Microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany), as described by Mitani et al (2000). The purity of CD14+ cells exceeded 95%, as determined using a FACScan flow cytometer (Becton Dickinson). The cytospin preparations of CD14+ cells were stained using non-specific esterase (α-naphthyl butyrate esterase) staining kits (Mutoh Chemical, Tokyo, Japan) according to the manufacturer's procedures. The CD14+ cells expressed non-specific esterase activity (data not shown). In some experiments, PBMCs were plated in 85-mm Falcon tissue culture dishes (Becton Dickinson Labware, Lincoln Park, USA) in 10% FCS RPMI-1640 medium at 37°C in a humidified atmosphere flushed with 5% CO2 for 2 h. Both adherent and non-adherent fractions were collected.

Culture of CD14+ cells CD14+ cells were cultured at a concentration of 5 × 105 cells/ml in a 24-well tissue culture plate (Nunc, Roskilde, Denmark) in RPMI-1640 medium supplemented with FCS, autologous plasma, autologous serum or HSA. GM-CSF and IL-4 were added on the initial day of culture. In some experiments, TNF-α was added consecutively to cultures. Cultures were incubated at 37°C in a humidified atmosphere flushed with 5% CO2 for a designated number of days. Half the culture media was replaced with fresh medium containing growth factors every 3–4 d. The numbers of viable cells were counted using trypan blue dye exclusion methods.

Phagocytosis assay DC (2 × 105) prepared from cultures with GM-CSF and IL-4 were incubated with 1 mg/ml FITC–dextran (40 000 mw; Sigma, St. Louis, USA) at 4°C or 37°C for 1 h in 10% FCS RPMI-1640 medium. FITC–dextran uptake was stopped by the addition of cold Ca2+-, Mg2+-free phosphate-buffered saline (PBS) containing 1% FCS, and the samples were washed four times with cold 1% FCS PBS. Uptake of FITC–dextran by the cells was analysed using a FACScan flow cytometer (Becton Dickinson).

Phenotypic analysis For single or double staining of cell surface molecules, the cells were processed for incubation with FITC-conjugated mAbs and/or PE-conjugated mAbs for 30 min on ice. The samples were washed three times with PBS and then analysed using a FACScan flow cytometer. The dead cells were gated out, depending on scatter properties. For suspension staining of an intracytoplasmic molecule, MPO, we used commercially available reagents (Cytofix/Cytoperm Kit, PharMingen, according to the manufacturer's instructions). Cells were fixed in formaldehyde-based fixation medium at 4°C for 20 min. After washing twice with PBS, pH 7·2, the cells were resuspended in permeabilization buffer containing sodium azide and saponin, and stained at 4°C for 45 min with mAb. After washing with PBS containing sodium azide, the cells were analysed using a FACScan flow cytometer.

Synthetic peptides and peptide-pulsing of DC A human immunodominant peptide, HER2p63 (TYLPTNASL), derived from the c-erbB-2/HER2/neu protooncogene was synthesized by Mitsubishi Chemical Co., Yokohama, Japan, as described previously by Nagata et al (1997) and Okugawa et al (2000). This peptide sequence has HLA-A2402 anchor motifs and the purity of peptides exceeded 95%. Peptides were dissolved in PBS, pH 7·4, at a concentration of 1 mg/ml and stored at 4°C until use. DC (1 × 106) were pulsed with HER2p63 at 10 µmol/l for 1 h at room temperature and for an additional 1 h at 37°C in a humidified atmosphere flushed with 5% CO2 (Ikuta et al, 2000; Okugawa et al, 2000). HER2p63-pulsed DC were assessed for enzyme-linked immunospot (ELISPOT) assay.

ELISPOT assay The ELISPOT assay was carried out as follows: 96-well nitrocellulose MAHA S4510 Millipore plates (Millipore, Bedford, USA.) were coated with 10 µg/ml of anti-interferon-γ (IFN-γ) mAb, 1-D1K (Mabtech, Stockholm, Sweden) overnight at 4°C. The plates were washed with PBS and treated with 10% heat-inactivated human AB serum for 2 h at 37°C to block non-specific binding. PBMCs (1 × 106 cells/well) from HLA-A2402+ volunteers were cultured with HER2p63-pulsed or non-pulsed irradiated DC in 24-well plates (Nunc) containing RPMI-1640 medium with 10% autologous serum for 7 d. After washing three times with PBS, the PBMCs were recultured with newly prepared HER2p63-pulsed or non-pulsed irradiated DC in RPMI-1640 medium with 10% autologous serum in the presence of 20 IU/ml of IL-2. After a 7 d culture period, the PBMCs were washed three times with PBS and added at designated concentrations to the ELISPOT plates together with 1 × 105 cells/ml of newly prepared HER2p63-pulsed or non-pulsed irradiated DC. After a 24 h incubation, the plates were washed six times with PBS containing 0·05% Tween, incubated with 1 µg/ml of biotinylated anti-IFN-γ mAb, 7-B6–1 (Mabtech), at 37°C for 2 h, and washed and reacted with 1 µg/ml of streptoavidin-alkaline phosphatase (Mabtech) for 1 h. After washing three times with PBS, the plates were stained using an alkaline phosphatase conjugate substrate Kit (BioRad, Hercules, USA). The spots were counted using the Axioplan 2 imaging analysis system (Carl Zeiss Vision, Hallbergmoos, Germany).

Results

Expression of CD14 and generation of DC by PBMCs

PBMCs were separated into four populations, based on adherence and CD14 expression. The adherent CD14+, adherent CD14, non-adherent CD14+ and non-adherent CD14 fractions were tested for DC generation using 5 d culture in 10% FCS RPMI-1640 medium with GM-CSF and IL-4. Previously, we had observed that when CD14+ cells were cultured in 10% FCS RPMI-1640 medium with GM-CSF and IL-4, CD1a+CD14 DC and CD1aCD14+ macrophages were generated (Mitani et al, 2000). Because of this, we evaluated CD1a+CD14 cells as DC under these conditions. The results from a representative study and are presented in Fig 1. CD14+ cells comprised approximately 15% and 10% in adherent cell and non-adherent cell fractions respectively. The majority of DC were recovered from CD14+ cells in both adherent and non-adherent cell fractions. In contrast, CD14 cells in adherent and non-adherent cell fractions gave rise to a small number of DC. CD14+ cells were then used as target cells for subsequent experiments.

Figure 1.

Generation of DC by adherent (Ad) CD14+, Ad CD14, non-adherent (Non-Ad) CD14+ and non-Ad CD14 fractions in PBMCs. PBMCs (40 × 106) were separated into Ad CD14+, Ad CD14, non-Ad CD14+ and non-Ad CD14 fractions. Cells in each fraction were cultured with GM-CSF plus IL-4 at a density of 5 × 105 cells/ml/well in 24-well plates. After a 5 d culture period,the cell number was determined. The cells were immunostained with anti-CD1a and anti-CD14 and CD1a+CD14 cells were evaluated as DC. Data are representative of five experiments.

Generation of DC by CD14+ cells supported by autologous plasma, autologous serum or HSA

In ex vivo generation of DC for clinical use, it is mandatory to omit FCS from cultures. Other investigators reported the use of autologous plasma or autologous serum in the ex vivo manipulation of DC for clinical trials (Murphy et al, 1996; Thurner et al, 1999; Titzer et al, 2000). Our preliminary experiments with HSA revealed the possibility that HSA could be substituted for autologous plasma and autologous serum. As human plasma and serum allow for generation of CD1a DC as well as CD1a+ DC, as confirmed by the expression of HLA-A,B,C, HLA-DR, CD80, CD86, CD40, CD54 and CD14 molecules (Duperrier et al, 2000; Pietschmann et al, 2000), we evaluated CD1a+CD14 and CD1aCD14 cells as DC obtained with cultures containing human plasma or serum. CD1a+CD14, but not CD1aCD14 cells, were observed in cultures with HSA. The profile of CD1a and CD14 expression of the cells in cultures with 1% human autologous plasma, 1% human autologous serum or 2% HSA are presented in Fig 2A. To search for the culture conditions that most efficiently support the differentiation of CD14+ cells into DC, we studied and compared the ability of autologous plasma, autologous serum and HSA to generate DC from CD14+ cells (Fig 2B). To define the optimal concentrations of autologous plasma, autologous serum and HSA, we added 0·1–10% of autologous plasma, autologous serum or HSA to cultures. When CD14+ cells from 10 healthy volunteers were cultured in the presence of autologous plasma, the highest number of DC was recovered at 1%, 2%, 5% and 10% in five, three, one and one cases respectively. With respect to 1%, 2% and 5% autologous serum, these gave rise to an optimal generation of DC in five, one and four cases respectively. When we added HSA to cultures in simultaneous experiments, 2% HSA was the most potent in generating DC in all cases tested. The number of DC obtained with 2% HSA was consistently higher than those obtained with autologous plasma or serum at all concentrations tested in each case. When all 10 cases were evaluated collectively, DC recovery with 2% HSA was significantly high compared with DC recovery in cultures conducted with respective concentrations of autologous plasma or autologous serum. The addition of 5% or 10% HSA to cultures reduced the generation of DC (data with 10% HSA not shown). We confirmed that this reduction was attributed to additives in HSA, sodium acetyltryptophan and sodium caprylate (data not shown).

Figure 2.

Generation of DC by CD14+ cells in cultures supplemented with autologous plasma, autologous sera and HSA. CD14+ cells were cultured with GM-CSF plus IL-4 at a density of 5 × 105 cells/ml/well in 24-well plates. After a 5 d culture period, the cells were harvested, enumerated and assessed for immunostaining using CD1a-FITC and CD14-PE. (A) Two-colour cytograms of the cells cultured with 1% autologous plasma, 1% autologous serum and 2% HSA in a representative case, case 6. In each protocol, cells in the indicated gates were enumerated as DC, as described in Materials and methods. (B) The numbers of DC generated with 0·1%, 1%, 2%, 5% and 10% autologous plasma or autologous sera, or 0·1%, 0·5%, 1%, 2% and 5% HSA in 10 cases. Data are presented as the mean of triplicate cultures. The difference between the number of DC generated with 2% HSA and each of the numbers with autologous plasma and autologous sera is significant at P < 0·01 using a paired t-test.

Effects of autologous plasma and autologous sera serially obtained on the generation of DC by CD14+ cells

Next, we tested whether plasma or sera samples obtained serially from the same volunteer can affect the generation of DC by CD14+ cells. Plasma and serum samples were obtained five times at more than 1 week intervals over a 6 month period from cases 3 and 5. To add autologous plasma and autologous serum to cultures at the concentration of 1%, we selected Cases 3 and 5, both of whom had the highest recovery of DC at 1% of plasma or serum (Fig 2B). The results are compared with those obtained with 2% HSA and are presented in Fig 3. Although the numbers of DC recovered varied considerably among different plasma or serum samples, 2% HSA generated the highest number of DC. We then determined if the batch of HSA would influence the generation of DC in our culture system and, for this, we used 20 batches from three different manufacturers. The generation of DC did not vary significantly among the individual batches of HSA (data not shown).

Figure 3.

Generation of DC by CD14+ cells in cultures supplemented with autologous plasma and autologous serum obtained at different time points or HSA. CD14+ cells from cases 3 and 5 were cultured with GM-CSF plus IL-4 at a density of 5 × 105 cells/ml/well in 24-well plates. After a 5 d culture period, the cells were harvested and the number of DC counted. Autologous plasma and autologous serum samples were obtained at five different time points with at least a 1 week interval for over 6 months. Autologous plasma, autologous serum and HSA were added to cultures at the concentrations of 1%, 1% and 2% respectively. Data are determined by assigning a value of 100% to the number of DC recovered in cultures with 10% FCS and presented as the mean of triplicate cultures. *Differences from the number of DC generated with 2% HSA is significant at P < 0·01 using a paired t-test. †Differences from the number of DC generated with 2% HSA is significant at P < 0·05 using a paired t-test.

Quantification of phagocytosis of DC generated in cultures with autologous plasma, autologous serum or HSA

To test the phagocytic function of DC generated in cultures with autologous plasma, autologous serum or HSA in the presence of GM-CSF and IL-4, we evaluated the temperature-dependent uptake of FITC–dextran by the DC using flow cytometry (Fig 4). The amounts of dextran particles incorporated were similar among DC generated in cultures with autologous plasma, autologous serum or HSA. Uptake of dextran was measurable at 37°C but not at 4°C. When the phagocytic activity was assayed with DC obtained from cultures in the presence of GM-CSF plus IL-4 for 5 d and GM-CSF plus IL-4 plus TNF-α for an additional 2 d, the level was reduced modestly compared with DC prepared with GM-CSF plus IL-4 (data not shown).

Figure 4.

Uptake of FITC–dextran by DC cultured with autologous plasma, autologous serum or HSA in the presence of GM-CSF and IL-4. Autologous plasma, autologous serum and HSA were added to cultures at a concentration of 1%, 1% and 2% respectively. The cells were incubated with FITC–dextran at 4°C or 37°C for 1 h, washed three times with cold PBS and analysed using a FACScan flow cytometer. For the analysis, we used total populations because CD14+ macrophages were not generated in cultures with autologous plasma, autologous serum or HSA in the presence of GM-CSF and IL-4, as shown in Fig 2A. The dotted and solid lines show the uptake of FITC–dextran at 4°C and 37°C respectively. Results are representative of four experiments.

Phenotype of DC generated in cultures with autologous plasma, autologous serum or HSA

HSA supported the generation of immature DC from CD14+ cells most effectively compared with autologous plasma or autologous sera obtained at various time points (Fig 3). We then investigated immunophenotypes of mature DC that were cultured with FCS, autologous plasma, autologous serum or HSA in the presence of GM-CSF and IL-4 for 5 d, followed by TNF-α for an additional 2 d. Analysis of surface markers of DC are presented in Fig 5. Cells obtained in different preparations expressed high levels of HLA-A,B,C, HLA-DR, CD80, CD86, CD40 and CD54, a finding which is consistent with features of mature DC. Similar to observations with GM-CSF and IL-4 (Fig 2A), DC cultured with HSA expressed CD1a, whereas CD14+ cells differentiated into CD1a+ and CD1a DC in the presence of autologous plasma or autologous serum.

Figure 5.

Phenotypic analysis of DC cultured with FCS, autologous plasma, autologous serum or HSA. CD14+ cells from case 6 were cultured at a density of 5 × 105 cells/ml/well in 24-well plates in the presence of GM-CSF plus IL-4 for 5 d, followed by TNF-α for an additional 2 d. FCS, autologous plasma, autologous serum and HSA were added to cultures at the concentrations of 10%, 1%, 1% and 2% respectively. After a 7 d culture period, the cells were harvested and processed for staining with a panel of mAbs, as indicated. Left panels show two-colour staining with CD1a-FITC and CD14-PE. Phenotypes with cells in the indicated gates in the left panels were analysed. The dotted and solid lines show the expression of isotype-matched controls and indicated molecules respectively. Results are representative of eight experiments.

CD8+ T-cell response induced by DC obtained from cultures with FCS, autologous plasma, autologous serum or HSA

To assess the ability of DC obtained from cultures with FCS, autologous plasma, autologous serum and HSA to prime antigen-specific CD8+ T cells, CD8+ T-cell responses induced by autologous DC pulsed with or without an immunodominant peptide HER2p63 (TYLPTNASL), which is derived from the c-erbB-2/HER2/neu protooncogene, were tested in PBMCs from two volunteers (cases 3 and 8) using an ELISPOT assay (Fig 6). As we wanted to add 1% autologous plasma and 1% autologous serum to cultures, cases 3 and 8 (for whom the optimal generation of DC was observed at 1% )(Fig 2B) were selected for this assay. A similar pattern for the number of HER2p63-specific IFN-γ-secreting cells was seen in both cases. Although the mean of HER2p63-specific IFN-γ-secreting cells induced by DC cultured with autologous serum or HSA was larger than that induced by DC cultured with FCS or autologous plasma, there was no significant difference in numbers of spots induced by DC generated in different conditions of cultures.

Figure 6.

ELISPOT assay with autologous DC pulsed with HER2p63 peptides. Autologous DC were prepared from two HLA-A2402+ volunteers, cases 3 and 8. CD14+ cells were cultured with FCS, autologous plasma, autologous serum or HSA at a density of 5 × 105 cells/ml/well in 24-well plates in the presence of GM-CSF plus IL-4 for 5 d and followed by TNF-α for an additional 2 d. FCS, autologous plasma, autologous serum and HSA were added to cultures at the concentrations of 10%, 1%, 1% and 2% respectively. After a 7 d culture period, the cells were harvested and processed for ELISPOT assay. Differences between the numbers of IFN-γ-specific spots obtained with HER2p63-pulsed and non-pulsed DC were evaluated as HER2-specific lymphocytes. Each bar represents the mean ± SD of numbers of spots for 2 × 105 PBMCs in triplicate wells. There were no significant differences among numbers of IFN-γ-specific spots obtained with DC in different preparations.

Discussion

Advances in our understanding of tumour immunity, such as the identification of molecular targets recognized by tumour-specific CTL, have encouraged new approaches for tumour-specific immunotherapy. As DC have the capacity to abrogate T-cell tolerance to tumour antigens (Gong et al, 1998), immunization of cancer patients with DC pulsed with tumour antigens or antigenetic peptides represent a promising candidate for cancer vaccines. For in vivo induction and elicitation of tumour-specific CTL responses using DC, several issues, including route of administration and delivery to the optimal sites of DC, remain to be determined. Our present study focused on how best to prepare DC.

The major sources of DC for ex vivo generation are CD34+ haemopoietic progenitor cells, precursor cells in PBMCs and monocytes in peripheral blood. The use of PBMCs, including monocytes, has an advantage for therapeutic applications over CD34+ haemopoietic progenitor cells because a large number of the target cells can be obtained easily. We first examined cytological properties of DC precursors in PBMCs in the context of adherence and CD14 expression. Although CD14+ cells accounted for a minor population of adherent and non-adherent cell fractions, the majority of DC emerged from CD14+ cells in both fractions. These findings indicate that DC precursors were numerous in a CD14+ population in PBMCs, irrespective of adherence. It is likely that the use of CD14+ cells as a source of DC precursors is feasible for generation of sizable numbers of pure DC.

The number of DC generated may be one of several major important issues for potential clinical use. In previous clinical studies, autologous plasma, autologous serum and FCS were used as serum or protein components to support the generation of DC from their precursors in cultures (Hsu et al, 1996; Murphy et al, 1996; Nestle et al, 1998; Lim & Bailey-Wood, 1999; Morse et al, 1999; Reichardt et al, 1999; Thurner et al, 1999; Mackensen et al, 2000; Titzer et al, 2000). The use of FCS in cultures to prepare DC precludes their practical application because FCS contains xenogenic proteins and other substances that may affect the function of DC. It is possible that the use of allogeneic human plasma or serum may lead to the transmission of infectious pathogens. These limitations suggest that autologous plasma or autologous serum would be a less hazardous choice for the generation of DC. However, our data from dose–response studies with 0·1–10% autologous plasma or autologous serum show that their optimal doses for the generation of DC vary considerably among individuals (Fig 2). Also, the numbers of DC recovered in cultures supplemented with autologous plasma or autologous serum obtained at different time points differed (Fig 3). These findings suggest that it is necessary to determine the appropriate concentration of autologous plasma or autologous serum for use in DC vaccine therapy. Investigators in several laboratories reported the generation of DC under serum-free conditions (Chen et al, 1998; Mitterer et al, 1998; Morse et al, 1999; Duperrier et al, 2000; Tarte et al, 2000). They used serum-free media supplemented with several human-derived proteins and were not fully chemically defined. Some of the proteins may affect functional properties of DC obtained in culture. We intended to search for a defined culture condition under which the generation of DC from CD14+ monocytes is reproducibly supported. After an extensive survey of culture conditions, we found that HSA could replace autologous plasma or autologous serum in cultures for generation of DC from CD14+ monocytes. The addition of 2% HSA unequivocally supported the generation of a maximal number of DC for all individuals tested. Moreover, the number of DC in cultures with 2% HSA was significantly higher compared with the numbers of DC obtained with various concentrations of autologous plasma or autologous sera. These observations imply that with the use of 2% HSA it is not necessary to define the optimal concentration using dose–response studies in respective patients before vaccination. In addition, there were no differences in DC generation among the batches of HSA that we tested. Nevertheless, it may be necessary to screen batches of HSA before clinical use.

We were also concerned with biological features of DC generated in HSA-containing cultures. DC generated in the presence of HSA were effective in phagocytic activity, similar to DC obtained from cultures with autologous plasma or autologous serum. In terms of the expression of MHC and co-stimulatory molecules at the cell surface, DC obtained from cultures with autologous plasma, autologous serum and HSA were similar. In agreement with previous reports (Bender et al, 1996; Chen et al, 1998; Duperrier et al, 2000; Pietschmann et al, 2000), our cultures with autologous plasma or autologous serum gave rise not only to CD1a+ DC but also to CD1a DC. However, DC obtained from cultures with HSA expressed CD1a molecules. Recently, it has been discovered that the CD1 molecule family can present lipid and glycolipid antigens to T cells by mechanisms that differ from those of the MHC molecules (Percelli & Modlin, 1999; Sugita & Brenner, 2000). Antigen presentation through the CD1 pathway might be exploited for lipid-based vaccines and adjuvants. We reported that HER2, derived from the c-erbB-2/HER2/neu protooncogene, was found to be a potent tumour rejection antigen in a murine in vivo experimental system, and we identified the immunodominant peptide, HER2p63 (TYLPTNASL), which has HLA-A2402 anchor motifs (Nagata et al, 1997; Okugawa et al, 2000). By using HER2p63, DC prepared in HSA-containing culture were tested for CTL priming potential. The results from ELISPOT assay indicate that our culture system supplemented with HSA can support the generation of DC with the potential to prime and induce HER2-specific CTL in a peptide-based manner. The ELISPOT assay enables the detection and quantification of peptide-specific T cells in the course of peptide-based vaccination protocols (Thurner et al, 1999; Titzer et al, 2000).This assay system may also be used to assess functions of DC prepared for immunotherapy and to detect the peptide-specific T-cell responses before vaccines. We are currently planning a DC-based vaccine trial with HER2p63 for HLA-A2402+ patients with HER2-expressing tumours, using DC prepared in cultures supplemented with HSA.

Acknowledgment

We thank M. Ohara for critical comments.

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