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Prepublished online as a Blood First Edition Paper on May 8, 2003; DOI 10.1182/blood-2002-12-3854.
Blood, 1 September 2003, Vol. 102, No. 5, pp. 1753-1763 Functional comparison of DCs generated in vivo with Flt3 ligand or in vitro from blood monocytes: differential regulation of function by specific classes of physiologic stimuliFrom the The Melbourne Tumour Biology Branch, Ludwig Institute for Cancer Research, Austin and Repatriation Medical Centre, Heidelberg, Victoria, Australia; Medizinische Klinik und Poliklinik V, University of Heidelberg, Heidelberg, Germany; and The Walter and Eliza Institute of Medical Research, Parkville, Victoria, Australia.
Dendritic cells (DCs) are a family of leukocytes that initiate T- and B-cell immunity against pathogens. Migration of antigen-loaded DCs from sites of infection into draining lymphoid tissues is fundamental to the priming of T-cell immune responses. In humans, the major peripheral blood DC (PBDC) types, CD1c+ DCs and interleukin 3 receptor–positive (IL-3R+) plasmacytoid DCs, are significantly expanded in vivo with the use of Flt3 ligand (FL). DC-like cells can also be generated from monocyte precursors (MoDCs). A detailed comparison of the functional potential of these types of DCs (in an autologous setting) has yet to be reported. Here, we compared the functional capacity of FL-expanded CD1c+ PBDCs with autologous MoDCs in response to 3 different classes of stimuli: (1) proinflammatory mediators, (2) soluble CD40 ligand trimer (CD40L), and (3) intact bacteria (Escherichia coli). Significant differences in functional capacities were found with respect to changes in phenotype, migratory capacity, cytokine secretion, and T-cell stimulation. MoDCs required specific stimuli for the expression of functions. They responded vigorously to CD40L or E coli, expressing cytokines known to regulate interferon-  (IFN-) in T cells (IL-12p70, IL-18, and IL-23), but
required prostaglandin E2 (PGE2) during stimulation to
migrate to chemokines. In contrast, PBDCs matured in response to minimal
stimulation, rapidly acquired migratory function in the absence of
PGE2-containing stimuli, and were low cytokine producers.
Interestingly, both types of DCs were equivalent with respect to stimulation
of allogeneic T-cell proliferation and presentation of peptides to cytotoxic T
lymphocyte (CTL) lines. These distinct differences are of particular
importance when considering the choice of DC types for clinical
applications.
Dendritic cells (DCs) are rare bone marrow–derived cells, involved in antigen capture, processing, and presentation. DCs are uniquely able to prime a naive T-cell response.1,2 Because of their critical role in orchestrating the immune response, there is increasing interest in using DCs as cellular vaccine adjuvants in the immunotherapy of cancer.3,4 A variety of soluble factors and pathogen signals are known to activate DCs.1,2 Thus, the maturation state of vaccine-loaded DCs will probably be critical for their regulation of appropriate T-cell immune responses. Three main sources of DCs have been used in clinical trials: DCs derived from (1) CD34+ progenitor cells, (2) CD14+ monocytes, and (3) peripheral blood DC precursors. Generation of CD34+-derived DCs requires 10 to 28 days in in vitro culture,5,6 while DCs generated in vitro from CD14+ monocytes (MoDCs) require 5 to 7 days.7,8 Although their physiologic relevance in vivo remains unclear, MoDCs are the major DC type used in vaccine-based clinical studies.3,4 MoDCs have also been used to establish many of the biologic paradigms of DC function.1,2 An alternative, and perhaps more physiological, source of DCs in humans is provided by the immature DC populations found in peripheral blood.9,10 At least 2 peripheral blood DC (PBDC) populations, constituting fewer than 1% of total mononuclear cells, exist in human peripheral blood: CD1c+ PBDCs and interleukin 3 receptor–positive (IL-3R+) plasmacytoid DCs (PDCs).9-13 Several cytokines are known to expand the number of these PBDC types in vivo, including granulocyte colony-stimulating factor (G-CSF) and Flt-3 ligand (FL).9,14,15 FL expands both human CD1c+ PBDCs and IL-3R+ PDCs9,10,14-18 and has antitumor effects in animal models.19-21 It has been suggested that the CD1c+ PBDC subset in peripheral blood is related to the CD14-derived dermal DCs and to germinal center DCs.6,22,23 Both of these types of DCs appear to be of myeloid origin and can differentiate into Langerhans cells in the presence of transforming growth factor– 
(TGF-).12,24
However, little is currently known of the functional differences between the
CD1c+ PBDC and MoDC types (eg, antigen uptake capacity, migration,
cytokine secretion, and regulation of T-cell function). Few direct comparisons of DC types have been reported. Comparisons of CD34+-derived DCs and MoDCs suggest that CD34+-derived DCs may be superior at activating low-frequency, peptide-specific cytotoxic T lymphocytes.25-28 Other studies have reported that IL-3R+ PDCs are functionally different from MoDCs.13,29-36 However, few of these studies have directly compared DC functions in an autologous setting; most have compared DC types among allogeneic donor sources. Thus, the degree to which donor variation contributes to the observed functional differences may be significant. We performed a clinical trial that evaluated FL (to expand PBDC numbers) with or without peptide vaccination in patients with malignant melanoma (M.J. Shackleton et al, submitted manuscript, 2003). The present study describes the functional analysis of FL-expanded CD1c+ PBDCs isolated from these patients and compares them with autologous MoDCs. Furthermore, CD1c+ PBDCs and autologous MoDCs from healthy donors were also compared to exclude the possibility that functional differences among DC types from cancer patients were due to the cancer itself or that alterations in DC behavior were due to FL administration. We found major differences between the responses of MoDCs and CD1c+ PBDCs toward 3 different classes of physiologic stimuli with respect to migratory function, cytokine production, and regulation of T-cell function.
DCs were cultured in RPMI 1640 (Trace Biosciences, Melbourne, Australia) supplemented with 20 mM HEPES (N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid), 60 mg/L penicillin G, 12.6 mg/L streptomycin, 2 mM L-glutamine, 1% nonessential amino acids, and 10% heat-inactivated fetal calf serum (FCS) (CSL, Melbourne, Australia) in a 5% CO2 incubator. Mixed leukocyte reactions (MLRs) were performed in Iscove modified Dulbecco medium (IMDM) (Gibco, Grand Island, NY) with 5% pooled normal human serum (gift from the Victorian Tissue Typing Service, Royal Melbourne Hospital, Melbourne, Australia) in a 10% CO2 incubator. Monoclonal antibodies and cytokines
Flow cytometric analysis of DCs and T cells was performed with the
following monoclonal antibodies: fluorescein isothiocyanate (FITC)–
conjugated immunoglobulin G1 (IgG1) isotype control; phycoerythrin
(PE)–conjugated IgG1 isotype control; anti-CD1a; anti-CD1c; anti-CD1d;
anti-CD45RA; anti-CD80; anti-CD83; anti-CD86; anti-CD123 (IL-3R Cell sources CD1c+ PBDCs and monocytes were isolated (1) from peripheral blood mononuclear cells (PBMCs) of patients with stage II, III, or IV melanoma enrolled in a phase 1 clinical study (LUD-97-012) (M.J. Shackleton et al, submitted manuscript, 2003) receiving 14 consecutive days of FL (Amgen) (25 µg/kg/d) alone or in combination with peptide vaccines or (2) from buffy packs from healthy donors provided by the Australian Red Cross Blood Bank (Southbank, Melbourne, Australia). In the present study, the various types of DCs were examined from patients with minimal residual disease to exclude the possible issues of advanced cancer on decreasing the functional capacity of DCs via the release of immunosuppressive cytokines. Blood for monocyte isolation was taken prior to administration of FL, and on day 15 for CD1c+ PBDC isolation. The Protocol Review Committee of the Ludwig Institute for Cancer Research and the Human Research Ethics Committee of the Austin and Repatriation Medical Centre (Heidelberg, Victoria, Australia) approved the protocol, and informed consent was obtained from all patients.
CD14+ monocytes were affinity purified by means of the MACS CD14
isolation kit (Miltenyi Biotech) and cultured (7 days) in RPMI/10% FCS (5
x 105/mL) with GM-CSF (40 ng/mL) and IL-4 (500 U/mL) in
24-well plates to generate MoDCs (more than 95% of cultured cells). On day 7,
all wells were pooled and readjusted to a DC concentration of 5 x
105/mL. Maturation-inducing factors were added on day 7, and cells
and supernatants were harvested on day 8 or 9 for functional assessment.
Cytokines and other stimuli in the present study (eg, TNF- Enrichment of CD1c+ PBDCs from FL-treated patients and healthy volunteers
CD1c+ PBDCs were enriched from frozen PBMC samples obtained from
the clinical trial (LUD-97-012) (M.J. Shackleton et al, submitted manuscript,
2003). After thawing, CD14+ monocytes, CD19+ B cells,
and CD3+ T cells were depleted by means of immunomagnetic beads
(MACS; Miltenyi Biotech) according to the manufacturer's instructions. This
depletion procedure routinely yielded greater than 60%
CD1c+CD14– HLA-DR+ PBDCs as assessed by
fluorescence-activated cell sorter (FACS). The enriched PBDCs were then
stained with anti-CD1-FITC (Biosource, Camarillo, CA), anti-CD123–PE
(IL-3R
Assays were performed as previously described.37 Briefly, lower chambers of Transwell plates (8.0-µm pore size) (Costar, Corning, NY) were filled with 500 µL RPMI/10% FCS with or without chemokines: CCL21 (macrophage inflammatory protein 3–
[MIP-3]) (300 ng/mL); CCL19 (6Ckine) (100 ng/mL); or CXCL12 (stromal
cell–derived factor 1-alpha [SDF-1 ; 30 ng/mL]) (all from
Peprotech). DCs (1 to 2 x 104) were added in 50 µL
RPMI/10% FCS into the upper chamber. After 2 hours, cells in the lower
chambers were harvested, concentrated to 50 µL volumes in Eppendorf tubes,
and counted microscopically with a hemocytometer. Each stimulation condition
was performed in triplicate wells. RNA isolation and cDNA synthesis
Total RNA was isolated from MoDCs and CD1c+ PBDCs by means of an
RNeasy Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer's
instructions. In brief, cells were lysed and homogenized in lysis buffer
containing guanidine isothiocyanate and Quantitative real-time PCR
Predeveloped assay reagents (PDARs) for IL-12p35, IL-12p40, and IL-18 were
obtained from Applied Biosystems and used in multiplex reactions with 18S rRNA
PDAR (Applied Biosystems) for normalization. Primers and probe for IL-23p19
were designed with the use of Primer Express software, version 1.5a (Applied
BioSystems). Gene expression levels were quantitated by means of ABI Prism
7700 Sequence Detection System (Applied Biosystems). PCR reactions were set up
in 96-well plates (25 µL per reaction) according to the manufacturer's
instructions and analyzed by means of the SDS program, version v1.7 (Applied
BioSystems). Relative expression was calculated by the Cytokine ELISAs
Cytokine secretion by stimulated DCs or by allogeneic T cells was measured
by cytokine enzyme-linked immunosorbent assays (ELISAs). Cytokine ELISA kits
were purchased for IL-2, IL-5, IL-6, IFN- T-cell purification and mixed leukocyte reaction
Allogeneic CD2+ T lymphocytes were obtained by rosetting PBMCs
with aminoethylisothiouronium (AET)–treated sheep red blood cells. T
cells were between 88% and 95% pure on the basis of CD3 staining. Varying
numbers of DCs were cultured in round-bottomed 96-well plates in triplicate
with 105 allogeneic PBMCs for 5 days in IMDM with 5% human serum.
After 5 days, 200 µL supernatants were harvested, and fresh medium
containing 1 µCi (0.037 MBq) [3H]thymidine (DuPont, Sydney, MA)
per well was added for 8 hours. Cells were transferred onto a glass fiber
filter (Wallac, Turku, Finland), and [3H]thymidine incorporation
was measured by means of an NXT TopCount Betaplate scintillation counter
(Packard, Meriden, CT). In separate experiments, the CD2+ T cells
(1 x 107) were labeled with 5-(and 6) carboxyfluorescein
diacetate succinimidyl ester (CFSE) (0.01 mM) in serum-free phosphate-buffered
saline (PBS) in the dark (10 minutes at room temperature. T cells were then
washed and cultured (3 x 105) with immature or mature MoDCs
(1 x 104) in round-bottomed 96-well plates in triplicate for
5 days. On day 5, cultures were restimulated with freshly matured MoDCs in the
presence of 10 µg/mL Brefeldin A at 37°C for 8 hours. Cells were
harvested, pelleted, and stained with anti-CD8–APC and
CD3–Cy-Chrome (BD Biosciences Pharmingen), washed again, and then fixed
with 1% paraformaldehyde (ProSciTech, Thuringowan, Australia)/PBS before
staining with FITC-conjugated anti–IFN- DC-peptide presentation to a cytotoxic T-lymphocyte (CTL) line
First, 6 x 10–6 to 6 x
10–12 M HLA-A2–restricted peptides NY-ESO-1b
(amino acids 157 through 165, sequence SLLMWITQC) (Biological Production
Facility, Ludwig Institute for Cancer Research, Heidelberg, Australia) and
Epstein-Barr virus (EBV) (BMLF1 sequence amino acids 280-288, GLCTLVAML,
Austin Research Institute, Melbourne, Australia) were treated at room
temperature for 1 hour with 500 µM Tris (tris(hydroxymethyl)aminomethane)
(2-carboxyethyl)–phosphine hydrochloride (TCEP) (Pierce, Rockford, IL)
in cystine-free Dulbecco modified Eagle medium (Cys-free DMEM) (Gibco) to
reduce dimerized peptides to monomeric form. MoDCs or CD1c+ PBDCs
or the transporter associated with antigen processing (TAP)–deficient T2
cells were resuspended in Cys-free DMEM, and equal volumes were added to the
reduced peptide and pulsed at room temperature for 30 minutes. The DCs or T2
cells were then washed once and resuspended in RPMI/10% FCS and 10 µg/mL
Brefeldin A at a cell concentration of 1 x 106/mL. Then, 100
µL peptide-pulsed DCs or T2 cells were incubated with 100 µL
peptide-specific T cells (APC-effector ratio of 1:1) at 37°C for 4 hours
in a 96-well U-bottom plate. Cells were pelleted, stained with anti-CD8
Cy-Chrome, washed, and then fixed with 1% paraformaldehyde (ProSciTech)/PBS
before staining with FITC-conjugated anti–IFN-
Cell morphology and culture of MoDCs and CD1c+ PBDCs CD1c+ PBDCs were purified from the PBMCs of melanoma patients (with minimal residual disease) treated with FL by removal of lineage-positive cells by monoclonal antibody (mAb)–magnetic-activated cell sorting (MACS) bead depletion and cell sorting of lineage-negative cells on the basis of CD1b/c and HLA-DR expression to greater than 97% purity. CD1c+ PBDCs showed poor viability if cultured in medium alone, but viabililty was substantially improved when they were cultured with GM-CSF and IL-4. FL-expanded CD1c+ PBDCs were morphologically identical to their counterparts from untreated individuals with a typical multilobulated nuclear morphology (Figure 1A). To avoid issues relating to the myelopoietic effects of FL upon monocyte development in vivo,9 the present study generated autologous MoDCs from CD14+ monocytes isolated from blood samples taken prior to FL administration. Immature MoDCs (GM-CSF plus IL-4) were morphologically distinct from freshly isolated CD1c+ PBDCs, being larger with round or kidney-shaped nuclear morphology and more extensive cytoplasm (Figure 1B). Both FL-expanded CD1c+ PBDCs and MoDCs displayed morphologic features typical of mature DCs following stimulation with CD40L, including prominent dendritic processes (Figure 1C-D).
Phenotypic analysis and maturation of MoDCs and CD1c+ PBDCs Freshly isolated CD1c+ PBDCs were phenotypically immature, expressing low levels of the maturation markers CD80, CD83 (Figure 2A), and CD86 (data not shown). Consistent with previous reports, CD1a was constitutively expressed on MoDCs but was not present on freshly isolated PBDCs (Figure 2A). In contrast, CD1c+ PBDCs, but not MoDCs, expressed CD1d. Both DC types expressed CD1c (Figure 2A) as well as CD1b, CD11c, CD13, CD33, and CD54 (data not shown), consistent with a putative myeloid origin. Whereas CD1c+ PBDCs spontaneously up-regulated the expression of CD80, CD83, and CD86 following overnight culture in medium containing GM-CSF and IL-4,38 MoDCs required maturation with specific combinations of stimuli.38 As previously reported, FL-expanded CD1c+ PBDCs showed heterogeneous expression of MMR.9 Additionally, discrete subpopulations within the CD1c+ PBDC gate were also detected on the basis of CCR6 and BDCA-3 expression (Figure 2B). The percentage expression for MMR, CCR6, and BDCA-3 suggests that multiple subpopulations are likely to exist. Interestingly, CD1c+ PBDCs up-regulated surface expression of CD83 (Figure 2C) and HLA-DR (data not shown) more rapidly than MoDCs, regardless of maturational stimulus. Furthermore, the mean fluorescence intensity of these markers was an order of magnitude greater for CD1c+ PBDCs compared with MoDCs (Figure 2C).
Induction of cytokine secretion by highly purified CD1c+ PBDCs and MoDCs DCs produce several types of cytokines following stimulation with pathogen or CD40L, such as IL-6, IL-10, and IL-12p70. We compared cytokine secretion by CD1c+ PBDCs and MoDCs in response to different classes of physiologic stimuli. Figure 3A shows that MoDCs secreted considerably more IL-6 compared with CD1c+ PBDCs, particularly in response to E coli. Similarly, MoDCs secreted IL-10 in response to a range of stimuli, but produced the highest levels of IL-10 following stimulation with E coli (Figure 3B). As previously reported, the addition of PGE2 to CD40L or to E coli decreased the amount of IL-10 produced by MoDCs.37
IL-12p70 is critical for the induction of IFN- Induction of cytokine secretion by CD1c+ PBDCs following initial in vitro culture prior to stimulation
Previous reports, as well as this study, indicate that freshly isolated
CD1c+ PBDCs are relatively poor producers of cytokines following
immediate
stimulation.37,38
However, 2 studies have shown that CD1c+ PBDCs can produce IL-12p70
following in vitro stimulation. In both studies, PBDCs were initially cultured
(thus matured) for at least 24 hours prior to stimulation with
lipopolysaccharide (LPS) or
CD40L.18,39
We, therefore, evaluated whether in vitro maturation of CD1c+ PBDCs
enhanced their responsiveness to IL-12p70–inducing stimuli such as
CD40L, intact E coli, or the combination of IL-1 Figure 3D also demonstrates that the low production of cytokines by CD1c+ PBDCs shown in Figure 3A-C was not due to the attenuating effects of IL-4 since similar levels of IL-12p70 were produced regardless of whether IL-4 was present or absent from the stimulation cocktail. Using quantitative real-time PCR (qRT-PCR), we examined whether the low levels of IL-12p70 produced by CD1c+ PBDCs are due to low levels of IL-12p35 or IL-12p40 mRNA expression. Figure 4A-B demonstrates that for MoDCs, the levels of IL-12p35 mRNA expression correlated with IL-12p70 production by ELISA. Similarly, for CD1c+ PBDCs, we found that low IL-12p70 secretion correlated with low expression of IL-12p35 and IL-12p40 mRNA (Figure 4A-B). Finally, the novel IL-23p19 mRNA was neither constitutively expressed by freshly isolated CD1c+ PBDCs nor induced following stimulation. In contrast, immature MoDCs constitutively expressed IL-23p19 mRNA, which was further increased following stimulation with E coli (Figure 4C).
IL-18, like IL-12p70 and IL-23, can induce IFN- Analysis of migratory capacity of MoDCs and CD1c+ PBDCs
Migration of antigen-loaded DCs toward lymphoid organs is critical for the
initiation of T-cell immunity and requires the expression of the chemokine
receptor CCR7 to respond to the lymph node–directing chemokines CCL19
(MIP-3
It has been proposed that MoDCs depend upon exogenous PGE2 as a consequence of IL-4's blocking endogenous PGE2 production by immature MoDCs.41 Alternatively, CD1c+ PBDCs, which are efficient migratory cells in the absence of PGE2-containing stimuli, may secrete higher levels of PGE2 in culture and thus not depend upon exogenous PGE2 to acquire migratory function. To address these possibilities, we examined the levels of PGE2 produced in culture SN by the 2 DC types. As shown in Figure 5E, MoDCs and CD1c+ PBDCs constitutively secreted comparable levels of PGE2 in vitro, and these levels were further increased following stimulation with E coli. Although not conclusive, these data argue that the differences in migratory capacity between MoDCs and CD1c+ PBDCs are not simply due to differences in the endogenous production of PGE2. Comparison of T-cell stimulatory capacity of MoDCs and CD1c+ PBDCs Mature DCs are the most efficient stimulators of naive T cells. We investigated the relative ability of differentially matured CD1c+ PBDCs or autologous MoDCs to stimulate the proliferation and cytokine secretion of alloreactive T cells in an MLR. CD1c+ PBDCs and autologous MoDCs were equally effective in stimulating allo–T-cell proliferation (Figure 6). However, MoDCs required prior activation with various physiologic stimuli to induce maximal T-cell proliferation. In this regard, immature MoDCs (GM-CSF plus IL-4) were, on a per cell basis, 10 to 100 times less efficient at inducing T-cell proliferation than mature MoDCs (Figure 6). In contrast, CD1c+ PBDCs induced T-cell proliferation equivalent to that seen with MoDCs irrespective of the stimulation conditions. This is consistent with the fact that CD1c+ PBDCs fully mature in culture without the need for further stimulation.
T-cell proliferation and cytokine secretion induced by MoDCs and/or CD1c+ PBDCs
Next, we assessed DC-mediated cytokine secretion by alloreactive T cells in
a separate series of experiments. Induction of IFN-
The ability of MoDCs and CD1c+ PBDCs to stimulate T-cell
cytokine secretion was also assessed by measuring IL-2, IL-5, and IFN-
Presentation of synthetic peptide to CTL lines by MoDCs and CD1c+ PBDCs
Finally, to assess antigen presentation to T cells, different populations
of DCs were used in a peptide-antigen (peptide-Ag) presentation assay. In this
assay, peptide-specific T cells were induced to produce IFN-
The clinical application of DCs requires a detailed understanding of their functional potential and how best to manipulate this for optimal vaccine delivery and immune induction. Both PBDCs and MoDCs are currently being evaluated in anticancer immunotherapy trials.3,4,42 This study provides the first detailed, direct comparison of these 2 DC populations by comparing autologous DC types under identical conditions. Both FACS-sorted CD1c+ PBDCs and highly purified MoDCs were isolated from melanoma patients (with minimal residual disease) participating in a clinical trial evaluating FL as a vaccine adjuvant (M.J. Shackleton et al, submitted manuscript, 2003). Both DC types were cultured in the same media, containing GM-CSF and IL-4 for optimal viability.7,29 A variety of cancers have been shown to affect the generation of functionally mature MoDCs.43,44 Indeed, we found that MoDCs and PBDCs from patients with later stage, metastatic disease receiving FL expressed reduced functional capacities such as the ability to mature in response to in vitro stimulation and the ability to stimulate T cells. Furthermore, some of these patients expressed significant monocytosis following FL treatment as well as elevated serum levels of proinflammatory cytokines such as IL-6 (M.J. Shackleton et al, submitted manuscript, 2003). However, MoDCs and CD1c+ PBDCs used in the present studies were specifically derived from patients with minimal residual disease, and these DCs were found to be functionally similar to their counterparts from healthy individuals37,38,45 (also found in data not shown). Several important findings were made in the present study. First, CD1c+ PBDCs and autologous MoDCs are phenotypically and functionally distinct DCs, differing in their migratory ability and their capacity to secrete specific cytokines, including IL-6, IL-10, and IL-12p70. Second, the function of these 2 DC subtypes was regulated by different types of soluble mediators. Finally, although these 2 DC types were equivalent at presenting peptides and T-cell stimulation, they induced different levels of T-cell cytokines.
MoDCs and CD1c+ PBDCs are frequently considered to be similar
cell
populations.13,29-36
Although, the phenotypes of these 2 distinct DC subtypes are similar (eg,
expression of CD4 and the myeloid markers CD11c, CD13, and CD33), there are
several markers that distinguish them. For instance, MoDCs express CD1a but
not CD1d, whereas CD1c+ PBDCs express CD1d but not CD1a. In
addition, while the majority of MoDCs express the pattern recognition receptor
MMR, only a subset (8% to 15%) of freshly isolated CD1c+ PBDCs
expressed MMR. In this regard, CD1c+ PBDCs appear to be
phenotypically heterogeneous, composed of distinct subsets expressing surface
Ags not expressed on immature MoDCs (eg, CCR6 and/or BDCA-3). The percentage
of CD1c+ PBDCs expressing MMR, CCR6, or BDCA-3 suggests that
multiple subpopulations are likely to exist. It is unclear, however, whether
these markers define distinct subsets or represent the same PBDC population at
different stages of maturation. We and others have noted that freshly isolated
FL-mobilized PBDCs are immature cells that mature rapidly (CD80+,
CD83+, CD86+) following
culture.10,16,38,46
The present study also indicates that this occurs with greater amplitude than
for MoDCs. Although MoDCs and CD1c+ PBDCs express a similar
repertoire of pathogen-recognition receptors (eg, MMR, DEC205 and Toll-like
receptors),30,47,48
MoDCs produce higher levels of IL-1
Several cytokines can induce IFN- | |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
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Abstract
Introduction
Ct method and is
expressed relative to a calibrator, in this case the GM-CSF/IL-4 DC control as
previously
described.
