Comparison of CD34 and Monocyte-Derived Dendritic Cells from Mobilized Peripheral Blood from Cancer Patients
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《干细胞学杂志》
ABSTRACT
Dendritic cells (DCs) are potent antigen-presenting cells that are integral to the initiation of T-cell immunity. Two cell types can be used as a source for generating DCs: monocytes and CD34+ stem cells. Despite many investigations characterizing DCs, none have performed a direct paired comparison of monocyte and stem cell–derived DCs. Therefore, it is unclear whether one cell source has particular advantages over the other, or whether inherent differences exist between the two populations. We undertook the following study to determine if there were any differences in DCs generated from monocytes or CD34+ cells from mobilized peripheral blood. DCs were generated by culturing the adherent cells (monocytes) in interleukin-4 and GM-CSF for 7 days, or by culturing nonadherent cells (CD34+) in the presence of GM-CSF and tumor necrosis factor alpha for 14 days. The resulting DCs were compared morphologically, phenotypically, functionally, and by yield. We could generate morphologically and phenotypically similar DCs. Differences were encountered when expression levels of some cell surface markers were examined (CD86, HLA-DR). There was no difference in how the DCs performed in a mixed lymphocyte reaction (p = .3). Further, no statistical difference was discovered when we examined cellular (DC) yield (p = .1); however, there was a significant difference when yield was normalized to the starting number of monocytes or CD34+ cells (p = .016). Together, these data demonstrate that differences do exist between monocyte-derived DCs and CD34-derived DCs from the same cellular product (apheresis) from the same individual.
INTRODUCTION
Dendritic cells (DCs) are potent antigen-presenting cells that can stimulate T cells. Traditionally these cells have been difficult to isolate due to their low concentration in peripheral blood (~0.01%) and the lack of a DC-specific marker [1]. Since the development of technologies that allow one to culture DCs in vitro, the possibility of exploiting these cells in a number of immunotherapeutic strategies has become a reality [2, 4]. Many strategies have been described that use these cells to help generate meaningful immune responses against a variety of malignancies. In recent years, a number of clinical trials using dendritic immunization have been undertaken [5, 7]. These investigations have employed both CD34 and monocyte-derived DCs, but it is still unclear if one cellular source is better than the other.
Both techniques generate large numbers of functional DCs. A number of studies have attempted to more directly compare monocyte-derived and CD34+ cell–derived DCs to try and understand these differences and determine the best source for DCs. Some studies suggest that a lower outgrowth of cells occurs when monocytes are used than with protocols that use CD34+ cells. Monocyte-derived and CD34+ cell–derived DCs have been described to have similar morphology, phenotype, antigen uptake, and presentation ability [8, 9]; however, some investigations suggest that monocyte-derived DCs have weaker expression of co-stimulatory molecules such as intercellular cell adhesion molecule–1, B7-1, and B7-2 [10, 11]. Differences in expression levels of co-stimulatory molecules may affect function, while for others, no significant differences were noted [12, 13]. Other surface markers can also vary. CD34-derived DCs have been reported to express chemokine (CC motif) receptor–6 (CCR6), while CD14-derived DCs did not. This was evidenced by differences in the cell’s associated chemotactic response [14, 15].
These two DC populations have also been reported by some to have comparable cytokine mRNA expression and expression of peptide antigen and can process and present antigen and induce cytotoxic T lymphocyte (CTL)–dependent interferon production [9, 12]. Others have shown that CD34+- and monocyte-derived DCs are not functionally equivalent with regard to activation of antigen-specific CTL [9, 11]. Monocyte-derived and CD34+-derived DCs are also described as being similar when generated from cancer patients [16]. Differences in the capacity to activate CD8+ T cells have been described [9, 11]. Further differences exist when it comes to generation. Monocyte-derived DCs have the advantage of being able to use unmobilized blood as a cellular source; however, on a per cell basis, mobilized blood produces more DCs than does unmobilized peripheral blood [17]. Generation of monocyte-derived DCs is also less time consuming, resulting in a pure population of DCs in 7 days as opposed to 12–14 days when culturing stem cells into DCs [4]. Since CD34+ cells are only found in small numbers in peripheral and cord blood, a system exploiting monocyte differentiation is attractive to researchers. Alternatively, in patients treated with high-dose chemotherapy and autologous stem cell transplantation, mobilized peripheral blood is available, which could serve as an alternative cellular source. In many circumstances patients fail to mobilize or have low CD34+ counts; in such situations, exploitation of monocytes for DC vaccination may be advantageous.
Despite these investigations, it is still unclear whether any differences between CD34+- and monocyte-derived DCs are significant and if these variations are attributable to the cell or to the cellular source. Conflicting information exists, making assessment difficult. Previous studies have not examined cells from the same cellular product from the same patient. Further, they have used minimal numbers of patients, and they often compare monocyte-derived DCs from normal healthy individuals to CD34+- derived DCs from those with malignancies [16]. Our previous work demonstrates that these are not equivalent populations [16].
To date, studies that have compared populations of CD34+- and CD14+-derived DCs have demonstrated some differences, but few of these investigations focused on patients with malignancies and none used paired well-controlled samples to truly detect if differences do exist, as well as the source of any differences. To determine the best cellular source for culturing DCs, we generated DCs from CD34+ cells and CD14+ cells from mobilized peripheral blood. DCs were then directly compared morphologically, phenotypically, and functionally and based on yield.
MATERIAL AND METHODS
Patients and Mobilization of Stem Cells
Apheresis product (after mobilizing chemotherapy) was obtained from 15 cancer patients (Table 1). Patients had previously signed an institutional review board–approved consent. Disease-mobilizing chemotherapy (Table 1) was followed 7 days later by rhG-CSF (recombinant human granulocyte colony-stimulating factor, Neupogen [Amgen, Thousand Oaks, CA]—300 μg for<70 kg body weight, 480 μg for 70–100 kg, or 600 μg for >100 kg) subcutaneously daily until apheresis was complete. Apheresis was usually performed on day 7 using a Cobe Spectra Cell separator (Cobe Laboratories, Arvada, CO, http://www.cobecv.com), as previously described [17, 18].
Collection of Apheresis Product
A sample (0.25–1.0 ml) of apheresis product was collected for this study. Peripheral blood mononuclear cells (PBMCs) were isolated from the product by centrifugation (x 800g for 20 minutes at room temperature) on a Ficoll-Hypaque density gradient (C-Six Diagnostics, Germantown, WI). Briefly, the apheresis product was diluted in 30 ml of Hanks’ buffered saline solution (HBSS) (Sigma Chemical Corp., St. Louis, http://www.sigma-aldrich.com) and underlaid with 10 ml of Ficoll Hypaque prior to centrifugation. PBMCs were harvested, then washed three times in HBSS and finally resuspended in RPMI (Roswell Park Memorial Institute) 1640 medium (Invitrogen, Burlington, ON, Canada http://www.invitrogen.com) containing 2 mM L-glutamine (Invitrogen), 100 U penicillin per ml (Invitrogen), 5 μg streptomycin per ml (Invitrogen), 0.2 μg of amphotericin B per ml (Invitrogen), 1 mM sodium pyruvate (Invitrogen), and 0.1 mM nonessential amino acids (Invitrogen). Cells were then counted on a hemacytometer. Viability was assessed using Trypan Blue exclusion.
Dendritic Cell Generation
A total of 2 x 107 PBMCs were plated per T25 flask (Corning, Acton, MA http://www.corning.com) in RPMI-1640 medium. Flasks were incubated at 370°C, 5% CO2 for 2 hours. After 2 hours, nonadherent cells were removed, collected, and counted. Adherent cells were then resuspended in 10 ml RPMI-1640 medium containing 1% human AB serum (Sigma), 800 U/ml GM-CSF (R&D Systems, Minneapolis, http://www.rndsystems.com) and 1000 U/ml IL-4 (interleukin–4) (R&D Systems). After 7 days, cells were harvested. To generate CD34-derived DCs, 2 x 107 of the original nonadherent cells were plated per T25 flask in RPMI-1640 medium containing 1% human AB serum (Sigma), 800 U/ml GM-CSF (R&D Systems), and 500 U/ ml tumor necrosis factor alpha (R&D Systems). After 12 days, cells were harvested and counted.
Yield
To determine yield, cells from one T25 flask were harvested and then washed in 1x HBSS without calcium chloride, magnesium chloride, or magnesium sulfate. Cells were then resuspended in RPMI-1640 medium and counted using a hemacytometer. Percentage of yield was determined based on the starting number of cells seeded into the flask or the percentage of CD14+ cells or CD34+ cells in this starting population (PBMCs or nonadherent cells). Viability was determined by Trypan Blue exclusion.
Cell Staining and Microscopy
Morphology was determined by performing a cytospin, staining the cells with Diff Quik?’ (Dade Behring, Dudinsen Switzerland, http://www.dadebehring.com), and examining cells by light microscopy.
Immunolabeling and Fluorescence-Activated Cell Sorter (FACS) Analysis
Initial PBMC populations from apheresis product, nonadherent cells, and cultured DCs were all immunolabeled. The following antibodies were used for surface labeling: anti-CD3 (SK7) fluorescein isothiocyanate (FITC), anti-CD14-FITC (MP9), anti-HLA-DR-FITC (L243), anti-CD11c (S-HCL-3) phycoerythrin (PE), anti-CD34-PE (8G12) (all antibodies from Becton, Dickinson, San Jose, CA http://www.bd.com, anti-CD83-PE (HB15a) (Immunotech, Marseille, France, http://www.sfrl.fr/qui/ad_immuno-tech.html), and anti-CD86-PE (FUN-1) (BD Biosciences Pharmingen, San Diego, CA, http://www.bdbiosciences.com). Analysis was performed using a FACScan flow cytometer (Becton, Dickinson). At least 1 x 104 events per sample were evaluated.
Mixed Lymphocyte Reaction (MLR)
PBMCs were obtained from major histocompatibility complex mismatched healthy volunteers. These cells were used as responders. PBMCs (2 x 105/well) were cultured in RPMI-1640 medium containing 5% human serum in the presence or absence of irradiated stimulator cells (DCs) at 1 x 104, or 1 x 103, cells/well in 96-well round-bottomed plates. After 4 days, cells were pulsed with 1 μCi of tritiated thymidine ([3H]TdR)(MPBiomedicals,http://www.mpbio.com). At 18 hours later, cells were harvested using a PHD cell harvester (Brandel, Gaithersburg, MD, http://www.brandel.com) and counted on a liquid scintillation counter. Data are presented as fold increase.
Statistics
Data are expressed as means ± standard errors of the means. Data were analyzed using Student’s t-test for paired variates of an analysis of variance. The statistical software Graph Pad Version 2.0 (PRISM, San Diego, http://www.prism-software.com) was used.
RESULTS
To generate DCs, the apheresis product was first divided into adherent and nonadherent fractions. To ensure that we were enriching for CD34+ cells in our nonadherent population, initial PBMCs and nonadherent cells were immunolabeled and examined for CD14 and CD34 expression (Table 2). Nonadherent cells expressed a significant increase in CD34 expression and a concomitant drop in CD14 expression, as expected.
Morphology
DCs have a characteristic morphology that is critical in identifying cells as DCs [19, 20]. To determine if the cellular source affected the morphology, DCs were cultured from CD34- or CD14-enriched cells from each apheresis product sample. Cells were stained and examined by light microscopy. Morphology of the DCs did not depend on the cellular source (CD34+ or CD14+) (Fig. 1A, B). Both cell sources produced large cells with an irregular shape. They had large ovoid or lobed nuclei and lacked a granular cytoplasm. Short "dendritic" cytoplasmic processes could be observed extending from the cell body, as previously described [17, 21]. These cultured DCs had features similar to those of peripheral blood DCs, as described in the literature [19, 20].
Phenotype
To determine if the phenotype of cultured DCs varied with cellular source, DCs were cultured from CD14- and CD34-enriched cells. The resulting DCs were then harvested, immunolabeled, and examined by flow cytometry. Loss of CD14 expression is a hallmark of DC generation [22]. In all instances of both CD14- and CD34-derived DCs, the resulting cells expressed low levels of T-cell, monocyte, and stem cell markers (Fig. 2A), as is expected for a pure population of DCs[2,23–25]. There was no difference in CD11c expression, but CD14+-derived DCs had significantly higher levels of B-7 (CD86, p = .0003) and HLA-DR (p = .03) (Fig. 2B). Low levels of CD83 were evident on both cell populations, as these cultured DCs are expected to have a relatively immature phenotype. For each population, viability and purity were both >90%, as examined by light microscopy.
Yield
To determine if any differences in yield existed between the two populations, a maximum of 2 x 107 cells were placed in each tissue culture flask. Total cellular yield was based on the number of cells initially put into each flask. There was a trend toward an increased yield from CD34+-derived cells, but it was not statistically significant (Fig. 3A ,p = .1). Significantly more DCs were generated from CD34+ cells than from monocytes, however, when yield was normalized for the starting number of relevant cells (p = .02) (CD14+ or CD34+) (Fig. 3B). Viability was similar between the two groups.
All DCs coexpressed CD86 and HLA-DR but did not coexpress CD3 and CD14 as had been expected.
Function
To determine if there were functional differences between them, the CD14+- and CD34+-derived DCs were tested in an MLR. We found that the DCs from both cellular sources behaved equivalently, causing a similar increase in cell number (Fig. 4).
DISCUSSION
The study reported here examined the generation of DCs from different cellular sources. To date, most studies, in which DCs were used therapeutically, used CD14+-derived DCs, but conflicting information exists as to whether this is the best source of DCs. Only by comparing equivalent cell sources can this question be answered. For our study we chose mobilized peripheral blood from cancer patients as a source of monocytes and CD34+ stem cells. This allowed us to directly compare CD34+- and monocyte-derived DCs in a population that is targeted for DC vaccination strategies and to determine if differences or advantages exist for either cell source. Our previous work has demonstrated that minor differences exist among DCs cultured from patients with a variety of malignancies and normal donors, but we did not compare DC-generation protocols [17].
Although a number of studies comparing CD14+- and CD34+-derived DCs exist, this is the first and only study undertaken in which a direct-paired comparison was made of these two cell populations in a group of cancer patients. Previous studies have produced inconsistent findings. Problems with many of these investigations are comparisons between very disparate groups (i.e., peripheral blood from healthy individuals for monocyte-derived DCs and mobilized blood from cancer patients for CD34+ DCs; bone marrow versus peripheral blood) [12, 16, 26]. Our previous studies suggested that monocyte-derived DCs from breast cancer patients behave different from other groups (normal donors, lymphoma patients) and are not representative of all malignancies but are a special group. Therefore, comparing these groups as sources of CD34+- and CD14+-derived DCs is not appropriate. Peripheral blood and mobilized blood are also not equivalent [17], suggesting that cellular source or initial product is important.
In many studies, CD34+ cells are isolated using antibody-based separation techniques, while CD14+ cells are isolated by plastic adherence. We sought to normalize the manipulation of cells and avoid artificially activating or engaging the cells with antibodies. By performing a paired study within one patient group, we have avoided many of these confounding factors that may explain observed differences between studies.
We assessed 15 cancer patients in this study. CD34+ and CD14+-derived DCs were generated from mobilized peripheral blood and assessed morphologically, phenotypically, functionally, and by yield. In a paired assessment, both protocols generated DCs that were morphologically identical: large cells with prominent round or ovoid nuclei. Both methods generated cells that expressed similar levels of CD3, CD83, CD11c, CD34, and CD14. Surprisingly, CD34+-derived DCs expressed significantly less CD86 and HLA-DR than did paired CD34+-derived DCs; however, both DC populations performed well in an MLR. When the percentage of yield of starting cell number was examined, CD34+-derived DCs tended to generate more DCs, but this was not statistically significant. When yield was normalized to the number of CD34+ or CD14+ cells in the starting population, CD34+-derived DCs generated significantly more DCs than did CD14+-derived DCs.
In our study CD14+- and CD34+-derived DCs were morphologically similar. Some differences in phenotype were observed between the two DC populations. All cultured DCs expressed low levels of CD14, a hallmark for monocyte differentiation into DCs [22, 27]. In both populations, low levels of CD3 and CD34 were also consistent with DC differentiation. Low levels of CD83 were evident on all cultured DCs, as they are expected to have a relatively immature phenotype. As expected, all DC populations expressed significant levels of HLA-DR, but CD14-derived DCs had significantly higher expression than CD34+-derived DCs. Differences in CD86 expression were also observed with CD14+-derived DCs, which had significantly higher levels than those in CD34+-derived DCs. Such differences have not been reported before in such cell populations. Peripheral blood DCs have been shown to have higher surface expression of HLA-DR and CD86 than bone marrow DCs [29]. When there were no differences observed in the phenotype and when CD34+-derived DCs had increased levels of costimulatory molecules, the studies were comparing cells derived from individuals with malignancies (often with breast cancer) with normal healthy individuals [11, 13, 16, 26]. Increases in costimulatory molecules on CD34+-derived DCs may contribute to their enhanced function [26]; any increases we observed did not seem to translate into differences in function.
When tested in an MLR, all DCs performed similarly. This is similar to a small study with a mix of healthy donors and lymphoma and myeloma patients in which no significant differences in MLR were observed [13]. CD34+-derived DCs from breast cancer patients [16] have been reported to have higher costimulatory molecule expression and superior performance in an MLR. Bone marrow–derived DCs have also been reported to perform better in an MLR [28]. Some investigators have reported that monocyte-derived DCs function better in an MLR [12], but other groups do not note such differences [9] or have found that CD34+-derived DCs are more potent [26], suggesting that patient health and cellular source are very important and that subtle differences in cytokine selection, concentration, and administration can affect function. As well, our results demonstrate that by removing important contributors of variation these cells are functionally equivalent.
Previously, we have shown that storage conditions can influence cellular yield [21] and mobilization [17]. Not surprisingly, we found that stem cells generated more DCs on a per cell basis than monocytes did. Since no IL-4 was used when CD34-derived DCs were generated into DCs, the contribution of CD14+ cells to the final DC outcome would be minimal. Preliminary experiments demonstrated that at least 50% of the adherent cells were CD14+ and approximately 2% were CD34+. When we used this minimum as representative of the CD14+ population, there was still a statistically significant difference between the CD14-derived and CD34-derived DC yield (p = .04). Here we observed a trend to increased cell number in CD34+-derived DCs (but not statistically significant) when examined based on starting number of total cells in each flask. Other investigations also demonstrate that CD34+ cells and bone marrow yield a greater number of DCs [13, 28]. This indicates that there may be an advantage to using CD34+-derived DCs if cell number is an issue for reaching immunization targets. Further, this may be especially beneficial in individuals with very high CD34 counts. If CD34+ cells are low, using CD14 cells may be a reasonable alternative.
Culture conditions and culture periods are different for CD14-derived DCs and CD34-derived DCs. Further, slight variations in conditions can affect morphology and phenotype but do not appear to influence cellular function [13, 30]. It has also been shown that identical culture conditions produce a higher yield of bone marrow–derived DCs than blood-derived DCs, but they require twice as long to develop [28]. Thus, the yield of CD34+-derived DCs might even be higher using different and prolonged culture conditions.
In summary, CD34+-derived DCs may be a more attractive source of DCs for a clinical vaccination protocol, since cellular yield is superior and costimulatory differences do not appear to translate into functional differences. Nevertheless, because of the similarities between the two cellular sources, both are good sources for generating DCs and, outside of yield, one is not superior to the other. Thus, cellular availability could be an important factor in determining which DC-generation protocol to use if all other issues are equal.
ACKNOWLEDGMENTS
This work was supported by a grant from Partners in Health, Alberta Cancer Board. We would like to acknowledge the apheresis unit at the Foothills Medical Centre for sample collection and the bone marrow transplant unit at the Tom Baker Cancer Centre for helping to recruit patients to this study.
REFERENCES
Voorhis WCV, Valinsky J, Hoffman J et al. Relative efficacy of human monocytes and dendritic cells as accessory cells for T cell replication. J Exp Med 1983;158:174–191.
Sallusto F, Lanzavecchia A. Efficient presentation of soluble antigen by cultured human dendritic cells is maintained by granulocyte/macrophage colony-stimulating factor plus interleukin 4 and down regulated by tumor necrosis factor a. J Exp Med 1994;179:1109–1118.
Caux C, Dezutter-Dambuyant C, Banchereau J. GM-CSF and TNF- cooperate in the generation of dendritic Langerhans cells. Nat Biotechnol 1992;360:258–261.
Syme RM, Gluck S. Effects of cytokines on the culture and differentiation of dendritic cells in vitro. J Hematother Stem Cell Res 2001;10:37–45.
Tjoa B, Erickson S, Barren R et al. In vitro propagated dendritic cells from prostate cancer patients as a component of prostate cancer immunotherapy. Prostate 1995;27:63–69.
Nestle FO, Alijagic S, Gilliet M et al. Vaccination of melanoma patients with peptide- or tumor lysate-pulsed dendritic cells. Nat Med 1989;4:328–332.
Celluzzi CM, Mayordomo JI, Storkus WJ et al. Peptide-pulsed dendritic cells induce antigen-specific CTL-mediated protective tumor immunity. J Exp Med 1996;183:283–287.
Herbst B, Kohler G, Mackensen A et al. CD34+ peripheral blood progenitor cell and monocyte derived dendritic cells: a comparative analysis. Br J Haematol 1997;99:490–499.
Ferlazzo G, Wesa A, Wei WZ et al. Dendritic cells generated either from CD34+ progenitor cells or from monocytes differ in their ability to activate antigen-specific CD8+ T cells. J Immunol 1999;163:3597–3604.
Siena S, Nicola MD, Mortarini R et al. Expansion of immunostimulatory dendritic cells from peripheral blood of patients with cancer. The Oncologist 1997;2:65–69.
Mortarini R, Anichini A, Nicola MD et al. Autologous dendritic cells derived from CD34+ progenitors and from monocytes are not functionally equivalent antigen-presenting cells in the induction of melan-A/Mart-1(27–35)-specific CTLs from peripheral blood lymphocytes of melanoma patients with low frequency of CTL precursors. Cancer Res 1997;57:5534–5541.
Triozzi PL, Aldrich W. Phenotypic and functional differences between human dendritic cells derived in vitro from hematopoietic progenitors and from monocyte/macrophages. J Leukoc Biol 1997;61:600–608.
Chen B, Stiff P, Sloan G et al. Replicative response, immunophenotype, and functional activity of monocyte-derived versus CD34+ derived dendritic cells following exposure to various expansion and maturational stimuli. Clin Immunol 2001;98:280–292.
Power CA, Church DJ, Meyer A et al. Cloning and characterization of a specific receptor for the novel CC chemokine MIP-3 alpha from lung dendritic cells. J Exp Med 1997;186:825–835.
Greaves DR, Wang W, Dairaghi DJ et al. CCR6, a CC chemokine receptor that interacts with macrophage inflammatory protein 3alpha and is very highly expressed in human dendritic cells. J Exp Med 1997;186:837–844.
Ferlazzo G, Klein I, Paliard X et al. Dendritic cells generated from CD34+ progenitor cells with flt3 ligand, c-kit ligand, GM-CSF, IL-4, and TNF-alpha are functional antigen presenting cells resembling mature monocyte-derived dendritic cells. J Immunother 2000;23:48–58.
Syme RM, Duggan P, Stewart D et al. Generation of dendritic cells ex vivo: differences in steady state versus mobilized blood from patients with breast cancer and lymphoma and normal donors. J Hematother Stem Cell Res 2001;10:621–630.
Stewart DA, Guo D, Morris D et al. Superior autologous blood stem cell mobilization from dose-intensive cyclophosphamide, etoposide, cisplatin plus G-CSF than from less intensive chemotherapy regimens. Bone Marrow Transplant 1999;23:111–117.[CrossRef][Medline]
Knight SC, Stagg A, Hill S et al. Development and function of dendritic cells in health and disease. J Invest Dermatol 1992;99:33S–38S.
Zhou L, Tedder TF. Human blood dendritic cells selectively express CD83, a member of the immunoglobulin super family. J Immunol 1995;154:3821–3835.
Syme R, Callaghan D, Duggan P et al. Storage of blood for in vitro generation of dendritic cells. Cytotherapy 2002;4:271–276.
Chapuis F, Rosenzwajg M, Yagello M et al. Differentiation of human dendritic cells from monocytes in vitro. Eur J Immunol 1997;27:431–441.
Romani N, Gruner S, Brang D et al. Proliferating dendritic cell progenitors in human blood. J Exp Med 1994;180:83–93.
Thurner B, Roder C, Dieckmann D et al. Generation of large numbers of fully mature and stable dendritic cells from leukaphersis products for clinical application. J Immunol Methods 1999;223:1–15.
Steinman RM. The dendritic cell system and its role in immunogenicity. Annu Rev Immunol 1991;9:271–296.
Meierhoff G, Krause SW, Andreesen R. Comparative analysis of dendritic cells derived from blood monocytes or CD34+ hematopoietic progenitor cells. Immunobiology 1998;198:501–513.
Syme R, Gluck S. Generation of dendritic cells: role of cytokines and potential clinical applications. Transfus Apheresis Sci 2001;24:117–124.
Bai L, Feuerer M, Beckhove P et al. Generation of dendritic cells from human bone marrow mononuclear cells: advantages for clinical application in comparison to peripheral blood monocyte derived cells. Int J Oncol 2002;20:247–253.
Raje N, Gong J, Chauhan D et al. Bone marrow and peripheral blood dendritic cells from patients with multiple myeloma are phenotypically and functionally normal despite the detection of Kaposi’s sarcoma herpesvirus gene sequences. Blood 1999;93:1487–1495.
Garderet L, Cao H, Salamero J et al. In vitro production of dendritic cells from human blood monocytes for therapeutic use. J Hematother Stem Cell Res 2001;10:553–567.(Rachel Symea,b, Rubinder )
Dendritic cells (DCs) are potent antigen-presenting cells that are integral to the initiation of T-cell immunity. Two cell types can be used as a source for generating DCs: monocytes and CD34+ stem cells. Despite many investigations characterizing DCs, none have performed a direct paired comparison of monocyte and stem cell–derived DCs. Therefore, it is unclear whether one cell source has particular advantages over the other, or whether inherent differences exist between the two populations. We undertook the following study to determine if there were any differences in DCs generated from monocytes or CD34+ cells from mobilized peripheral blood. DCs were generated by culturing the adherent cells (monocytes) in interleukin-4 and GM-CSF for 7 days, or by culturing nonadherent cells (CD34+) in the presence of GM-CSF and tumor necrosis factor alpha for 14 days. The resulting DCs were compared morphologically, phenotypically, functionally, and by yield. We could generate morphologically and phenotypically similar DCs. Differences were encountered when expression levels of some cell surface markers were examined (CD86, HLA-DR). There was no difference in how the DCs performed in a mixed lymphocyte reaction (p = .3). Further, no statistical difference was discovered when we examined cellular (DC) yield (p = .1); however, there was a significant difference when yield was normalized to the starting number of monocytes or CD34+ cells (p = .016). Together, these data demonstrate that differences do exist between monocyte-derived DCs and CD34-derived DCs from the same cellular product (apheresis) from the same individual.
INTRODUCTION
Dendritic cells (DCs) are potent antigen-presenting cells that can stimulate T cells. Traditionally these cells have been difficult to isolate due to their low concentration in peripheral blood (~0.01%) and the lack of a DC-specific marker [1]. Since the development of technologies that allow one to culture DCs in vitro, the possibility of exploiting these cells in a number of immunotherapeutic strategies has become a reality [2, 4]. Many strategies have been described that use these cells to help generate meaningful immune responses against a variety of malignancies. In recent years, a number of clinical trials using dendritic immunization have been undertaken [5, 7]. These investigations have employed both CD34 and monocyte-derived DCs, but it is still unclear if one cellular source is better than the other.
Both techniques generate large numbers of functional DCs. A number of studies have attempted to more directly compare monocyte-derived and CD34+ cell–derived DCs to try and understand these differences and determine the best source for DCs. Some studies suggest that a lower outgrowth of cells occurs when monocytes are used than with protocols that use CD34+ cells. Monocyte-derived and CD34+ cell–derived DCs have been described to have similar morphology, phenotype, antigen uptake, and presentation ability [8, 9]; however, some investigations suggest that monocyte-derived DCs have weaker expression of co-stimulatory molecules such as intercellular cell adhesion molecule–1, B7-1, and B7-2 [10, 11]. Differences in expression levels of co-stimulatory molecules may affect function, while for others, no significant differences were noted [12, 13]. Other surface markers can also vary. CD34-derived DCs have been reported to express chemokine (CC motif) receptor–6 (CCR6), while CD14-derived DCs did not. This was evidenced by differences in the cell’s associated chemotactic response [14, 15].
These two DC populations have also been reported by some to have comparable cytokine mRNA expression and expression of peptide antigen and can process and present antigen and induce cytotoxic T lymphocyte (CTL)–dependent interferon production [9, 12]. Others have shown that CD34+- and monocyte-derived DCs are not functionally equivalent with regard to activation of antigen-specific CTL [9, 11]. Monocyte-derived and CD34+-derived DCs are also described as being similar when generated from cancer patients [16]. Differences in the capacity to activate CD8+ T cells have been described [9, 11]. Further differences exist when it comes to generation. Monocyte-derived DCs have the advantage of being able to use unmobilized blood as a cellular source; however, on a per cell basis, mobilized blood produces more DCs than does unmobilized peripheral blood [17]. Generation of monocyte-derived DCs is also less time consuming, resulting in a pure population of DCs in 7 days as opposed to 12–14 days when culturing stem cells into DCs [4]. Since CD34+ cells are only found in small numbers in peripheral and cord blood, a system exploiting monocyte differentiation is attractive to researchers. Alternatively, in patients treated with high-dose chemotherapy and autologous stem cell transplantation, mobilized peripheral blood is available, which could serve as an alternative cellular source. In many circumstances patients fail to mobilize or have low CD34+ counts; in such situations, exploitation of monocytes for DC vaccination may be advantageous.
Despite these investigations, it is still unclear whether any differences between CD34+- and monocyte-derived DCs are significant and if these variations are attributable to the cell or to the cellular source. Conflicting information exists, making assessment difficult. Previous studies have not examined cells from the same cellular product from the same patient. Further, they have used minimal numbers of patients, and they often compare monocyte-derived DCs from normal healthy individuals to CD34+- derived DCs from those with malignancies [16]. Our previous work demonstrates that these are not equivalent populations [16].
To date, studies that have compared populations of CD34+- and CD14+-derived DCs have demonstrated some differences, but few of these investigations focused on patients with malignancies and none used paired well-controlled samples to truly detect if differences do exist, as well as the source of any differences. To determine the best cellular source for culturing DCs, we generated DCs from CD34+ cells and CD14+ cells from mobilized peripheral blood. DCs were then directly compared morphologically, phenotypically, and functionally and based on yield.
MATERIAL AND METHODS
Patients and Mobilization of Stem Cells
Apheresis product (after mobilizing chemotherapy) was obtained from 15 cancer patients (Table 1). Patients had previously signed an institutional review board–approved consent. Disease-mobilizing chemotherapy (Table 1) was followed 7 days later by rhG-CSF (recombinant human granulocyte colony-stimulating factor, Neupogen [Amgen, Thousand Oaks, CA]—300 μg for<70 kg body weight, 480 μg for 70–100 kg, or 600 μg for >100 kg) subcutaneously daily until apheresis was complete. Apheresis was usually performed on day 7 using a Cobe Spectra Cell separator (Cobe Laboratories, Arvada, CO, http://www.cobecv.com), as previously described [17, 18].
Collection of Apheresis Product
A sample (0.25–1.0 ml) of apheresis product was collected for this study. Peripheral blood mononuclear cells (PBMCs) were isolated from the product by centrifugation (x 800g for 20 minutes at room temperature) on a Ficoll-Hypaque density gradient (C-Six Diagnostics, Germantown, WI). Briefly, the apheresis product was diluted in 30 ml of Hanks’ buffered saline solution (HBSS) (Sigma Chemical Corp., St. Louis, http://www.sigma-aldrich.com) and underlaid with 10 ml of Ficoll Hypaque prior to centrifugation. PBMCs were harvested, then washed three times in HBSS and finally resuspended in RPMI (Roswell Park Memorial Institute) 1640 medium (Invitrogen, Burlington, ON, Canada http://www.invitrogen.com) containing 2 mM L-glutamine (Invitrogen), 100 U penicillin per ml (Invitrogen), 5 μg streptomycin per ml (Invitrogen), 0.2 μg of amphotericin B per ml (Invitrogen), 1 mM sodium pyruvate (Invitrogen), and 0.1 mM nonessential amino acids (Invitrogen). Cells were then counted on a hemacytometer. Viability was assessed using Trypan Blue exclusion.
Dendritic Cell Generation
A total of 2 x 107 PBMCs were plated per T25 flask (Corning, Acton, MA http://www.corning.com) in RPMI-1640 medium. Flasks were incubated at 370°C, 5% CO2 for 2 hours. After 2 hours, nonadherent cells were removed, collected, and counted. Adherent cells were then resuspended in 10 ml RPMI-1640 medium containing 1% human AB serum (Sigma), 800 U/ml GM-CSF (R&D Systems, Minneapolis, http://www.rndsystems.com) and 1000 U/ml IL-4 (interleukin–4) (R&D Systems). After 7 days, cells were harvested. To generate CD34-derived DCs, 2 x 107 of the original nonadherent cells were plated per T25 flask in RPMI-1640 medium containing 1% human AB serum (Sigma), 800 U/ml GM-CSF (R&D Systems), and 500 U/ ml tumor necrosis factor alpha (R&D Systems). After 12 days, cells were harvested and counted.
Yield
To determine yield, cells from one T25 flask were harvested and then washed in 1x HBSS without calcium chloride, magnesium chloride, or magnesium sulfate. Cells were then resuspended in RPMI-1640 medium and counted using a hemacytometer. Percentage of yield was determined based on the starting number of cells seeded into the flask or the percentage of CD14+ cells or CD34+ cells in this starting population (PBMCs or nonadherent cells). Viability was determined by Trypan Blue exclusion.
Cell Staining and Microscopy
Morphology was determined by performing a cytospin, staining the cells with Diff Quik?’ (Dade Behring, Dudinsen Switzerland, http://www.dadebehring.com), and examining cells by light microscopy.
Immunolabeling and Fluorescence-Activated Cell Sorter (FACS) Analysis
Initial PBMC populations from apheresis product, nonadherent cells, and cultured DCs were all immunolabeled. The following antibodies were used for surface labeling: anti-CD3 (SK7) fluorescein isothiocyanate (FITC), anti-CD14-FITC (MP9), anti-HLA-DR-FITC (L243), anti-CD11c (S-HCL-3) phycoerythrin (PE), anti-CD34-PE (8G12) (all antibodies from Becton, Dickinson, San Jose, CA http://www.bd.com, anti-CD83-PE (HB15a) (Immunotech, Marseille, France, http://www.sfrl.fr/qui/ad_immuno-tech.html), and anti-CD86-PE (FUN-1) (BD Biosciences Pharmingen, San Diego, CA, http://www.bdbiosciences.com). Analysis was performed using a FACScan flow cytometer (Becton, Dickinson). At least 1 x 104 events per sample were evaluated.
Mixed Lymphocyte Reaction (MLR)
PBMCs were obtained from major histocompatibility complex mismatched healthy volunteers. These cells were used as responders. PBMCs (2 x 105/well) were cultured in RPMI-1640 medium containing 5% human serum in the presence or absence of irradiated stimulator cells (DCs) at 1 x 104, or 1 x 103, cells/well in 96-well round-bottomed plates. After 4 days, cells were pulsed with 1 μCi of tritiated thymidine ([3H]TdR)(MPBiomedicals,http://www.mpbio.com). At 18 hours later, cells were harvested using a PHD cell harvester (Brandel, Gaithersburg, MD, http://www.brandel.com) and counted on a liquid scintillation counter. Data are presented as fold increase.
Statistics
Data are expressed as means ± standard errors of the means. Data were analyzed using Student’s t-test for paired variates of an analysis of variance. The statistical software Graph Pad Version 2.0 (PRISM, San Diego, http://www.prism-software.com) was used.
RESULTS
To generate DCs, the apheresis product was first divided into adherent and nonadherent fractions. To ensure that we were enriching for CD34+ cells in our nonadherent population, initial PBMCs and nonadherent cells were immunolabeled and examined for CD14 and CD34 expression (Table 2). Nonadherent cells expressed a significant increase in CD34 expression and a concomitant drop in CD14 expression, as expected.
Morphology
DCs have a characteristic morphology that is critical in identifying cells as DCs [19, 20]. To determine if the cellular source affected the morphology, DCs were cultured from CD34- or CD14-enriched cells from each apheresis product sample. Cells were stained and examined by light microscopy. Morphology of the DCs did not depend on the cellular source (CD34+ or CD14+) (Fig. 1A, B). Both cell sources produced large cells with an irregular shape. They had large ovoid or lobed nuclei and lacked a granular cytoplasm. Short "dendritic" cytoplasmic processes could be observed extending from the cell body, as previously described [17, 21]. These cultured DCs had features similar to those of peripheral blood DCs, as described in the literature [19, 20].
Phenotype
To determine if the phenotype of cultured DCs varied with cellular source, DCs were cultured from CD14- and CD34-enriched cells. The resulting DCs were then harvested, immunolabeled, and examined by flow cytometry. Loss of CD14 expression is a hallmark of DC generation [22]. In all instances of both CD14- and CD34-derived DCs, the resulting cells expressed low levels of T-cell, monocyte, and stem cell markers (Fig. 2A), as is expected for a pure population of DCs[2,23–25]. There was no difference in CD11c expression, but CD14+-derived DCs had significantly higher levels of B-7 (CD86, p = .0003) and HLA-DR (p = .03) (Fig. 2B). Low levels of CD83 were evident on both cell populations, as these cultured DCs are expected to have a relatively immature phenotype. For each population, viability and purity were both >90%, as examined by light microscopy.
Yield
To determine if any differences in yield existed between the two populations, a maximum of 2 x 107 cells were placed in each tissue culture flask. Total cellular yield was based on the number of cells initially put into each flask. There was a trend toward an increased yield from CD34+-derived cells, but it was not statistically significant (Fig. 3A ,p = .1). Significantly more DCs were generated from CD34+ cells than from monocytes, however, when yield was normalized for the starting number of relevant cells (p = .02) (CD14+ or CD34+) (Fig. 3B). Viability was similar between the two groups.
All DCs coexpressed CD86 and HLA-DR but did not coexpress CD3 and CD14 as had been expected.
Function
To determine if there were functional differences between them, the CD14+- and CD34+-derived DCs were tested in an MLR. We found that the DCs from both cellular sources behaved equivalently, causing a similar increase in cell number (Fig. 4).
DISCUSSION
The study reported here examined the generation of DCs from different cellular sources. To date, most studies, in which DCs were used therapeutically, used CD14+-derived DCs, but conflicting information exists as to whether this is the best source of DCs. Only by comparing equivalent cell sources can this question be answered. For our study we chose mobilized peripheral blood from cancer patients as a source of monocytes and CD34+ stem cells. This allowed us to directly compare CD34+- and monocyte-derived DCs in a population that is targeted for DC vaccination strategies and to determine if differences or advantages exist for either cell source. Our previous work has demonstrated that minor differences exist among DCs cultured from patients with a variety of malignancies and normal donors, but we did not compare DC-generation protocols [17].
Although a number of studies comparing CD14+- and CD34+-derived DCs exist, this is the first and only study undertaken in which a direct-paired comparison was made of these two cell populations in a group of cancer patients. Previous studies have produced inconsistent findings. Problems with many of these investigations are comparisons between very disparate groups (i.e., peripheral blood from healthy individuals for monocyte-derived DCs and mobilized blood from cancer patients for CD34+ DCs; bone marrow versus peripheral blood) [12, 16, 26]. Our previous studies suggested that monocyte-derived DCs from breast cancer patients behave different from other groups (normal donors, lymphoma patients) and are not representative of all malignancies but are a special group. Therefore, comparing these groups as sources of CD34+- and CD14+-derived DCs is not appropriate. Peripheral blood and mobilized blood are also not equivalent [17], suggesting that cellular source or initial product is important.
In many studies, CD34+ cells are isolated using antibody-based separation techniques, while CD14+ cells are isolated by plastic adherence. We sought to normalize the manipulation of cells and avoid artificially activating or engaging the cells with antibodies. By performing a paired study within one patient group, we have avoided many of these confounding factors that may explain observed differences between studies.
We assessed 15 cancer patients in this study. CD34+ and CD14+-derived DCs were generated from mobilized peripheral blood and assessed morphologically, phenotypically, functionally, and by yield. In a paired assessment, both protocols generated DCs that were morphologically identical: large cells with prominent round or ovoid nuclei. Both methods generated cells that expressed similar levels of CD3, CD83, CD11c, CD34, and CD14. Surprisingly, CD34+-derived DCs expressed significantly less CD86 and HLA-DR than did paired CD34+-derived DCs; however, both DC populations performed well in an MLR. When the percentage of yield of starting cell number was examined, CD34+-derived DCs tended to generate more DCs, but this was not statistically significant. When yield was normalized to the number of CD34+ or CD14+ cells in the starting population, CD34+-derived DCs generated significantly more DCs than did CD14+-derived DCs.
In our study CD14+- and CD34+-derived DCs were morphologically similar. Some differences in phenotype were observed between the two DC populations. All cultured DCs expressed low levels of CD14, a hallmark for monocyte differentiation into DCs [22, 27]. In both populations, low levels of CD3 and CD34 were also consistent with DC differentiation. Low levels of CD83 were evident on all cultured DCs, as they are expected to have a relatively immature phenotype. As expected, all DC populations expressed significant levels of HLA-DR, but CD14-derived DCs had significantly higher expression than CD34+-derived DCs. Differences in CD86 expression were also observed with CD14+-derived DCs, which had significantly higher levels than those in CD34+-derived DCs. Such differences have not been reported before in such cell populations. Peripheral blood DCs have been shown to have higher surface expression of HLA-DR and CD86 than bone marrow DCs [29]. When there were no differences observed in the phenotype and when CD34+-derived DCs had increased levels of costimulatory molecules, the studies were comparing cells derived from individuals with malignancies (often with breast cancer) with normal healthy individuals [11, 13, 16, 26]. Increases in costimulatory molecules on CD34+-derived DCs may contribute to their enhanced function [26]; any increases we observed did not seem to translate into differences in function.
When tested in an MLR, all DCs performed similarly. This is similar to a small study with a mix of healthy donors and lymphoma and myeloma patients in which no significant differences in MLR were observed [13]. CD34+-derived DCs from breast cancer patients [16] have been reported to have higher costimulatory molecule expression and superior performance in an MLR. Bone marrow–derived DCs have also been reported to perform better in an MLR [28]. Some investigators have reported that monocyte-derived DCs function better in an MLR [12], but other groups do not note such differences [9] or have found that CD34+-derived DCs are more potent [26], suggesting that patient health and cellular source are very important and that subtle differences in cytokine selection, concentration, and administration can affect function. As well, our results demonstrate that by removing important contributors of variation these cells are functionally equivalent.
Previously, we have shown that storage conditions can influence cellular yield [21] and mobilization [17]. Not surprisingly, we found that stem cells generated more DCs on a per cell basis than monocytes did. Since no IL-4 was used when CD34-derived DCs were generated into DCs, the contribution of CD14+ cells to the final DC outcome would be minimal. Preliminary experiments demonstrated that at least 50% of the adherent cells were CD14+ and approximately 2% were CD34+. When we used this minimum as representative of the CD14+ population, there was still a statistically significant difference between the CD14-derived and CD34-derived DC yield (p = .04). Here we observed a trend to increased cell number in CD34+-derived DCs (but not statistically significant) when examined based on starting number of total cells in each flask. Other investigations also demonstrate that CD34+ cells and bone marrow yield a greater number of DCs [13, 28]. This indicates that there may be an advantage to using CD34+-derived DCs if cell number is an issue for reaching immunization targets. Further, this may be especially beneficial in individuals with very high CD34 counts. If CD34+ cells are low, using CD14 cells may be a reasonable alternative.
Culture conditions and culture periods are different for CD14-derived DCs and CD34-derived DCs. Further, slight variations in conditions can affect morphology and phenotype but do not appear to influence cellular function [13, 30]. It has also been shown that identical culture conditions produce a higher yield of bone marrow–derived DCs than blood-derived DCs, but they require twice as long to develop [28]. Thus, the yield of CD34+-derived DCs might even be higher using different and prolonged culture conditions.
In summary, CD34+-derived DCs may be a more attractive source of DCs for a clinical vaccination protocol, since cellular yield is superior and costimulatory differences do not appear to translate into functional differences. Nevertheless, because of the similarities between the two cellular sources, both are good sources for generating DCs and, outside of yield, one is not superior to the other. Thus, cellular availability could be an important factor in determining which DC-generation protocol to use if all other issues are equal.
ACKNOWLEDGMENTS
This work was supported by a grant from Partners in Health, Alberta Cancer Board. We would like to acknowledge the apheresis unit at the Foothills Medical Centre for sample collection and the bone marrow transplant unit at the Tom Baker Cancer Centre for helping to recruit patients to this study.
REFERENCES
Voorhis WCV, Valinsky J, Hoffman J et al. Relative efficacy of human monocytes and dendritic cells as accessory cells for T cell replication. J Exp Med 1983;158:174–191.
Sallusto F, Lanzavecchia A. Efficient presentation of soluble antigen by cultured human dendritic cells is maintained by granulocyte/macrophage colony-stimulating factor plus interleukin 4 and down regulated by tumor necrosis factor a. J Exp Med 1994;179:1109–1118.
Caux C, Dezutter-Dambuyant C, Banchereau J. GM-CSF and TNF- cooperate in the generation of dendritic Langerhans cells. Nat Biotechnol 1992;360:258–261.
Syme RM, Gluck S. Effects of cytokines on the culture and differentiation of dendritic cells in vitro. J Hematother Stem Cell Res 2001;10:37–45.
Tjoa B, Erickson S, Barren R et al. In vitro propagated dendritic cells from prostate cancer patients as a component of prostate cancer immunotherapy. Prostate 1995;27:63–69.
Nestle FO, Alijagic S, Gilliet M et al. Vaccination of melanoma patients with peptide- or tumor lysate-pulsed dendritic cells. Nat Med 1989;4:328–332.
Celluzzi CM, Mayordomo JI, Storkus WJ et al. Peptide-pulsed dendritic cells induce antigen-specific CTL-mediated protective tumor immunity. J Exp Med 1996;183:283–287.
Herbst B, Kohler G, Mackensen A et al. CD34+ peripheral blood progenitor cell and monocyte derived dendritic cells: a comparative analysis. Br J Haematol 1997;99:490–499.
Ferlazzo G, Wesa A, Wei WZ et al. Dendritic cells generated either from CD34+ progenitor cells or from monocytes differ in their ability to activate antigen-specific CD8+ T cells. J Immunol 1999;163:3597–3604.
Siena S, Nicola MD, Mortarini R et al. Expansion of immunostimulatory dendritic cells from peripheral blood of patients with cancer. The Oncologist 1997;2:65–69.
Mortarini R, Anichini A, Nicola MD et al. Autologous dendritic cells derived from CD34+ progenitors and from monocytes are not functionally equivalent antigen-presenting cells in the induction of melan-A/Mart-1(27–35)-specific CTLs from peripheral blood lymphocytes of melanoma patients with low frequency of CTL precursors. Cancer Res 1997;57:5534–5541.
Triozzi PL, Aldrich W. Phenotypic and functional differences between human dendritic cells derived in vitro from hematopoietic progenitors and from monocyte/macrophages. J Leukoc Biol 1997;61:600–608.
Chen B, Stiff P, Sloan G et al. Replicative response, immunophenotype, and functional activity of monocyte-derived versus CD34+ derived dendritic cells following exposure to various expansion and maturational stimuli. Clin Immunol 2001;98:280–292.
Power CA, Church DJ, Meyer A et al. Cloning and characterization of a specific receptor for the novel CC chemokine MIP-3 alpha from lung dendritic cells. J Exp Med 1997;186:825–835.
Greaves DR, Wang W, Dairaghi DJ et al. CCR6, a CC chemokine receptor that interacts with macrophage inflammatory protein 3alpha and is very highly expressed in human dendritic cells. J Exp Med 1997;186:837–844.
Ferlazzo G, Klein I, Paliard X et al. Dendritic cells generated from CD34+ progenitor cells with flt3 ligand, c-kit ligand, GM-CSF, IL-4, and TNF-alpha are functional antigen presenting cells resembling mature monocyte-derived dendritic cells. J Immunother 2000;23:48–58.
Syme RM, Duggan P, Stewart D et al. Generation of dendritic cells ex vivo: differences in steady state versus mobilized blood from patients with breast cancer and lymphoma and normal donors. J Hematother Stem Cell Res 2001;10:621–630.
Stewart DA, Guo D, Morris D et al. Superior autologous blood stem cell mobilization from dose-intensive cyclophosphamide, etoposide, cisplatin plus G-CSF than from less intensive chemotherapy regimens. Bone Marrow Transplant 1999;23:111–117.[CrossRef][Medline]
Knight SC, Stagg A, Hill S et al. Development and function of dendritic cells in health and disease. J Invest Dermatol 1992;99:33S–38S.
Zhou L, Tedder TF. Human blood dendritic cells selectively express CD83, a member of the immunoglobulin super family. J Immunol 1995;154:3821–3835.
Syme R, Callaghan D, Duggan P et al. Storage of blood for in vitro generation of dendritic cells. Cytotherapy 2002;4:271–276.
Chapuis F, Rosenzwajg M, Yagello M et al. Differentiation of human dendritic cells from monocytes in vitro. Eur J Immunol 1997;27:431–441.
Romani N, Gruner S, Brang D et al. Proliferating dendritic cell progenitors in human blood. J Exp Med 1994;180:83–93.
Thurner B, Roder C, Dieckmann D et al. Generation of large numbers of fully mature and stable dendritic cells from leukaphersis products for clinical application. J Immunol Methods 1999;223:1–15.
Steinman RM. The dendritic cell system and its role in immunogenicity. Annu Rev Immunol 1991;9:271–296.
Meierhoff G, Krause SW, Andreesen R. Comparative analysis of dendritic cells derived from blood monocytes or CD34+ hematopoietic progenitor cells. Immunobiology 1998;198:501–513.
Syme R, Gluck S. Generation of dendritic cells: role of cytokines and potential clinical applications. Transfus Apheresis Sci 2001;24:117–124.
Bai L, Feuerer M, Beckhove P et al. Generation of dendritic cells from human bone marrow mononuclear cells: advantages for clinical application in comparison to peripheral blood monocyte derived cells. Int J Oncol 2002;20:247–253.
Raje N, Gong J, Chauhan D et al. Bone marrow and peripheral blood dendritic cells from patients with multiple myeloma are phenotypically and functionally normal despite the detection of Kaposi’s sarcoma herpesvirus gene sequences. Blood 1999;93:1487–1495.
Garderet L, Cao H, Salamero J et al. In vitro production of dendritic cells from human blood monocytes for therapeutic use. J Hematother Stem Cell Res 2001;10:553–567.(Rachel Symea,b, Rubinder )