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*
Laboratory of Molecular Immunoregulation, Division of Basic Sciences, and
Intramural Research Support Program, Science Applications International Corp.-Frederick, National Cancer Institute-Frederick Cancer Research and Development Center, National Institutes of Health, Frederick, MD 21702-1201; and
Laboratory of Immunology, National Institute of Allergy and Infectious Disease, National Institutes of Health, Bethesda, MD 20892
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The trafficking pattern of DCs, like that of other leukocytes, is presumably controlled by many mediators. However, the direction of DC migration is primarily determined by chemotactic gradients formed by a variety of chemotactic factors, including classical chemoattractants and chemokines (3, 5, 10, 11, 12, 13, 14, 15, 16). Classical chemoattractants include formyl peptides (e.g., cleavage products of bacterial and mitochondrial proteins such as fMLP) (5, 10), products of host complement activation (e.g., C5a) (10), and lipid metabolites (e.g., platelet activating factor) (15), whereas chemokines consist of a superfamily of >30 structurally related proteins classified into the CXC, CC, CX3C, and C chemokine subfamilies (16, 17, 18). The effects of classical chemoattractants and chemokines are mediated by members of the G protein-coupled seven-transmembrane domain receptor superfamily (18, 19, 20). The reason that iDCs and mDCs are able to migrate toward different chemotactic factors is due to the fact that they express different sets of chemotactic receptors. So far, human iDCs have been shown to express CXCR1 (12, 21), CXCR2 (12), CXCR4 (12, 21, 22, 23, 24), CCR1 (12, 21, 22, 24), CCR2 (12, 21, 22), CCR3 (22, 24), CCR4 (21, 22), CCR5 (12, 21, 22, 23, 24), CCR6 (14, 25, 26, 27, 28, 29), CCR8,6 and probably CCR9 (30, 31, 32), whereas mDCs express only CXCR4 (21, 23) and CCR7 (14, 21, 33, 34, 35).
Studies of DC responses to chemoattractants have shown that human iDCs derived from either peripheral blood monocytes or cord blood CD34+ cells respond chemotactically to and express the receptor for platelet activating factor (15). DCs isolated from rat respiratory tract tissue or generated in vitro from human peripheral blood monocytes can be chemoattracted by fMLP and C5a (5, 10). Human skin-derived Langerhans’ cells, a prototype of iDCs, express the receptor for C5a (C5aR) and respond chemotactically to C5a (36). However, the regulation of DC responsiveness to fMLP and C5a and expression of formyl peptide receptor (FPR), the high-affinity receptor for fMLP, and C5aR upon DC maturation, have not been fully elucidated. Here, we investigated this issue by using human and murine DCs generated in vitro from either hemopoietic progenitor cells (HPC) or CD14+ peripheral blood monocytes. The results show that iDCs of both species express FPR and C5aR and are thus able to respond to fMLP and C5a. In contrast, mDCs differentially down-regulate FPR, but not C5aR, expression at mRNA and protein levels, resulting in a selective retention of responsiveness to C5a as compared with fMLP.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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IMDM was purchased from Life Technologies (Rockville, MD). RPMI
1640 was purchased from BioWhittaker (Walkersville, MD). Recombinant
human (rh) stromal cell-derived factor-1
(SDF-1), TNF-,
GM-CSF, IL-4, Flt3-ligand, stem cell factor, and thrombopoietin were
purchased from PeproTech (Rocky Hill, NJ). Recombinant murine (rm)
GM-CSF, IL-4, and TNF- were purchased from Biosource International
(Camarillo, CA), R&D Systems (Minneapolis, MN), and PharMingen (San
Diego, CA), respectively. fMLP, rhC5a, and chemicals unless otherwise
specified were purchased from Sigma (St. Louis, MO). FBS was purchased
from HyClone (Logan, UT). FITC-conjugated mouse anti-human C5aR
(CD88) was purchased from Serotec (Oxford, U.K.). Mouse anti-human
CD40 agonistic Ab (IgG1, 
, capable of stimulating B cell
proliferation in the presence of IL-4) was purchased from PharMingen.
Anti-CD83 was purchased from Coulter-Immunotech (Marseille, France).
The other Abs used for flow cytometry were purchased from PharMingen.
[3H]fMLP with a specific radioactivity of 2000
Ci/mmol and [3H]TdR with a specific
radioactivity of 2 Ci/mmol were purchased from NEN (Boston, MA). PCR
primers for C5aR were purchased from Stratagene (La Jolla,
CA).
Cell isolation and purification
Human PBMC were isolated by routine Ficoll-Hypaque density
gradient centrifugation. Monocytes were purified (>95%) from human
PBMC with the use of a MACS CD14 monocyte isolation kit (Miltenyi
Biotech, Auburn, CA) according to the manufacturer’s recommendation.
Cord blood CD34+ HPC (>95%) were purchased from
Poietics (Gaithersburg, MD). The DC precursors were amplified from
CD34+ HPC exactly as described (37)
by culturing the cells at 5 x 104 cells/ml
in IMDM supplemented with 20% FBS, 10-5 M DTT,
25 ng/ml rhFlt3-ligand, 10 ng/ml rh thrombopoietin, and 20 ng/ml rh
stem cell factor for 4 wk. The amplified DC precursors were
cryopreserved in IMDM containing 20% FBS and 10% DMSO until later
usage. Murine HPC were prepared from the bone marrow of C57BL/6 mice
(female, 7 wk) as described (38). Briefly, bone marrow
cells flushed from femur and tibia were depleted of RBC by ammonium
chloride treatment. For depletion of lymphocytes and Ia-positive cells,
the remaining cells were incubated with a mixture of mAb for 1 h
at 4°C followed by depletion with immunomagnetic beads coated with
anti-rat IgG (Dynal, Great Neck, NY). The mAbs used were
anti-B220/CD45R (PharMingen), anti-MHC class II (M5/114.15.2
anti-I-Ab, d, q and I-Ed,
k; American Type Culture Collection, Manassas, VA), and
anti-CD90 (Thy1; PharMingen). The resulting cells (
90%
Sca-1+/Lin-) were used as
murine HPC.
DC preparation
To generate human monocyte-derived iDCs, purified monocytes were
incubated at 1 x 106/ml in RPMI 1640 medium
(RPMI 1640 plus 10% FBS, 2 mM glutamine, 25 mM HEPES, 100 U/ml
penicillin, 100 µg/ml streptomycin) in the presence of rhGM-CSF (50
ng/ml) and rhIL-4 (1050 ng/ml) at 37°C in a
CO2 (5%) incubator for 7 days. For the
generation of human CD34+-derived iDCs, DC
precursors amplified from CD34+ HPC were
incubated at 5 x 105/ml in RPMI 1640 medium
in the presence of rhGM-CSF (50 ng/ml) and rhIL-4 (50 ng/ml) at 37°C
in a CO2 (5%) incubator for 2 days
(37). Murine iDCs were generated by the incubation of
murine HPC at 1 x 106/ml in RPMI 1640
medium in the presence of rmGM-CSF (50 U/ml) and rmIL-4 (10 ng/ml) at
37°C in a CO2 (5%) incubator for 5 days
(38). All of the cultures were fed with the same
cytokine-containing medium every 2–3 days. To induce DC maturation,
iDCs were cultured in the same cytokine mixtures with added TNF- (50
ng/ml) or anti-CD40 Ab (100 µg/ml) for 48 h at 37°C in a
CO2 (5%) incubator (21, 23, 27, 28).
Chemotaxis assay
DC migration was assessed using a 48-well microchemotaxis chamber technique as previously described (27, 39). Briefly, different concentrations of chemotactic factors were placed in the wells of the lower compartment of the chamber (Neuroprobe, Cabin John, MD), and DCs (106 cells/ml) were added to the wells of the upper compartment. The lower and upper compartments were separated by a 5-µm polycarbonate filter (Osmonics, Livermore, CA). After incubation at 37°C in humidified air with 5% CO2 for 1.5 h, the filters were removed and stained, and the cells migrating across the filter were counted with the use of a Bioquant semiautomatic counting system. The results are presented as number of cells per high power field.
Calcium mobilization
DCs (107 cells/ml in RPMI 1640 containing 10% FBS) were loaded with dye by incubating with 5 µM fura-2 (Molecular Probes, Eugene, OR) at 24°C for 30 min in the dark. Subsequently, the loaded cells were washed and resuspended (106 cells/ml) in saline buffer (138 mM NaCl, 6 mM KCl, 1 mM CaCl2, 10 mM HEPES, 5 mM glucose, and 1% BSA, pH 7.4). Each 2 ml of loaded DC suspension was then transferred into a quartz cuvette, which was placed in a luminescence spectrometer LS50 B (Perkin-Elmer, Beaconsfield, U.K). Ca2+ mobilization of the cells was measured by recording the ratio of fluorescence emitted at 510 nm after sequential excitation at 340 and 380 nm in response to chemotactic factors at various concentrations.
FACS
DCs were first washed three times with FACS buffer (PBS, 1% FBS, 0.02% NaN3, pH 7.4) and then stained with FITC-conjugated anti-CD88 or control Ab at room temperature for 30 min as recommended by the manufacturer. After washing three times with PBS, the stained DCs were fixed with 1% paraformaldehyde in PBS, stored at 4°C overnight, and analyzed the next day with a flow cytometer (Coulter Epics, Miami, FL).
MLR
Allogeneic MLR was performed as described (11). Briefly, purified allogeneic T cells (105/well) were cultured with different numbers of iDCs or mDCs in a 96-well flat-bottom plate for 7 days at 37°C in humidified air with 5% CO2. The proliferative response of T cells was examined by pulsing the culture with [3H]TdR (0.5 µCi/well) for another 18 h before harvesting. [3H]TdR incorporation was measured with a microbeta counter (Wallac, Gaithersburg, MD).
Binding assay
Equilibrium binding was performed in triplicate by adding a constant amount of [3H]fMLP and increasing amounts of unlabeled fMLP to individual 1.5-ml microfuge tubes, each containing 2 x 106 DCs suspended in RPMI 1640 containing 1% BSA, 2.5 mM HEPES, 0.05% NaN3. After incubation at 24°C with constant mixing for 20 min, the cells were extensively washed with cold PBS, and the cell-associated radioactivity was measured with a microbeta counter (Wallac).
RNA isolation, RT-PCR, and Northern blot
Total RNA from DCs was isolated by the use of TRIzol Reagent (Life Technologies). The RNAs were cleaned by treatment with RNase-free DNase I (Stratagene). RT-PCR was performed by the use of GeneAmp RNA PCR Kit (Roche Molecular Systems, Branchburg, NJ). Briefly, 100 ng of RNAs was used in the RT-PCR. After reverse transcription, C5a and GAPDH cDNA fragments were amplified by 30 cycles of PCR (denatured at 95°C for 30 s, annealing at 54°C for 30 s, and extension at 72°C for 1 min) with the last extension being performed at 72°C for 10 min. The primers for human C5aR were 5'-CCCAAGCTTGGGGCGGGGAATCAATGAATTTCAGCGA-3' and 5'-CCGCTCGAGCGGCTATCACATAGTGAAGGAGGACGCA-3'. The primers for human GAPDH were 5'-GATGACATCAAGAAGGTGGTGAA-3' and 5'-GTCTTACTCCTTGGAGGCCATGT-3'. PCR products were identified on 1–2% agarose gel after ethidium bromide staining and photodocumented. Northern blot was performed as described elsewhere (40) with minor modification. Briefly, total RNA (20 µg/lane) was fractionated on 1% agarose-formaldehyde gel and transferred to a nitrocellulose filter. The specific mRNA on the filter was detected by hybridization with a 32P-labeled cDNA probe at 42°C overnight in buffer comprising 50% formamide, 5x SSPE (1x SSPE is 0.15 M NaCl, 10 mM NaH2PO4, 10 mM EDTA, pH 7.4), 5x Denhardt’s solution, 1% SDS, and 100 µg/ml denatured salmon sperm DNA. Two cDNA probes were used: a 1000-bp HindIII-EcoRI fragment of human FPR cDNA and a 400-bp ß-actin cDNA fragment (41). The probes were labeled by a RadPrime DNA labeling kit (Life Technologies). After hybridization, the filters were washed with 2x SSC (1x SSC is 0.15 M NaCl, 15 mM sodium citrate, pH 7.0) containing 0.2% SDS and 0.1x SSC containing 0.1% SSC until a reasonably low background was obtained. The filter was dried and autoradiographed overnight at -80°C using a Kodak x-ray film (Rochester, NY).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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To address the effect of maturation of DCs on their response to
fMLP and C5a, we compared the chemotaxis and Ca2+
mobilization of iDCs and mDCs derived from human monocytes in response
to fMLP and C5a. iDCs migrated to both fMLP and C5a with bell-shaped
dose-response curves with optimal concentrations at
10-8 M and 10-9 M,
respectively (Fig. 1
, A and
C), confirming the previous reports (5, 10).
Both fMLP and C5a also induced intracellular Ca2+
mobilization by iDCs in a dose-dependent manner (Fig. 2, A and B,
upper panels). After treatment of iDCs with rhTNF- for
48 h to induce maturation, mDCs lost their responsiveness to fMLP
as measured by chemotaxis (Fig. 1B) and
Ca2+ mobilization (Fig. 2A,
lower panel), but maintained responsiveness to C5a in terms
of both chemotaxis (Fig. 1D) and Ca2+
mobilization (Fig. 2B, lower panel). Furthermore,
C5a was equally potent and efficacious for iDCs and mDCs because 1) it
induced the migration of similar numbers of iDCs and mDCs at identical
optimal concentration (10-9 M) under similar
experimental conditions (Fig. 1, C and D), and 2)
it mobilized intracellular Ca2+ to a similar
extent in concentrations ranging from 10-11 to
10-7 M (Fig. 2B). The result that
SDF-1 was chemotactic for both iDCs and mDCs (Fig. 1) is in
accordance with previous reports (21, 27).
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, iDCs were
CD1a+, CD11a+,
CD14-, CD40low,
CD83-, CD86low, and
HLA-DRmedium, whereas mDCs were
CD1a+, CD11a+,
CD14-, CD40high,
CD83+, CD86high, and
HLA-DRhigh. In addition, iDCs were unable to
stimulate allogeneic MLR whereas mDCs stimulated marked proliferation
of allogeneic T cells, especially at DC:T ratios equal to 1:100 or
1:10, as detected by [3H]TdR incorporation
(Table II). These data confirmed that
monocyte-derived iDCs and mDCs used in this study had the
characteristics of iDCs and mDCs.
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In vivo, down-regulation of the response of DCs to fMLP may result
from either down-regulation of FPR expression or homologous
desensitization due to the presence of an excessive amount of formyl
peptides in the environment, which can derive from either microbes or
disrupted mitochondria (19, 42, 43). Although homologous
desensitization is less likely to occur when DCs are induced to mature
in an in vitro culture system, we examined the expression of FPR and
C5aR by iDCs and mDCs to rule out this possibility. To this end, iDCs
and mDCs were tested for binding to [3H]fMLP.
iDCs specifically bound [3H]fMLP, and this was
inhibited in a dose-dependent manner by unlabeled fMLP, whereas mDCs
did not bind [3H]fMLP at all, suggesting that
mDCs greatly decrease their surface FPR expression (Fig. 3A). In contrast, both iDCs
and mDCs expressed comparable amounts of C5aR on their surfaces as
determined by FACS analysis after staining of the cells with
FITC-conjugated anti-CD88 (Fig. 3B). To determine
whether TNF- induces differential regulation of FPR and C5aR
expression at the transcriptional or posttranscriptional level, the
expression of FPR and C5aR mRNAs by iDCs and mDCs was further
investigated. By Northern blot analysis, iDCs were shown to express FPR
mRNA (Fig. 4A, lane
1), whereas FPR mRNA expression by mDCs was undetectable (Fig. 4A, lane 2). Reprobing of the same filter with a
32P-labeled actin cDNA fragment after stripping
yielded bands of nearly identical intensity, confirming that equal
amounts of RNAs were loaded in both lanes 1 and 2
(Fig. 4A). As expected, both iDCs and mDCs expressed a
similar level of C5aR mRNA as measured by RT-PCR using C5aR-specific
primers (Fig. 4B, lanes 1 and 2).
These results indicate that DCs, upon maturation induced by TNF-,
down-regulate their FPR expression while maintaining their C5aR
expression.
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DC maturation can be induced in vitro by treatment of iDCs with a
variety of agents including bacterial products (LPS, etc.) (14, 21, 23, 33), proinflammatory cytokines (TNF-, IL-1, etc.)
(14, 21, 23, 27, 28, 33), synthetic nucleic acids
(CpG-oligodeoxynucleotides, poly(I:C), etc.) (44, 45), and
macrophage-conditioned medium (34, 45), by cross-linking
of membrane CD43 (46) or by CD40 ligation with CD40 ligand
or anti-CD40 Ab (14, 21, 33). However, DC maturation
induced by TNF- and many other agents is considered reversible and
only that induced by poly(I:C), macrophage-conditioned medium, or
CD40 ligation is stable (21, 45, 47, 48). To determine
whether stable maturation of DCs affects the regulation of FPR and C5aR
expression, we prepared human mDCs by treatment of monocyte-derived
iDCs with anti-CD40 agonistic Ab. mDCs generated in this manner
showed phenotypic (CD40high,
CD83+, CD86high, and
HLA-DRhigh) and functional (capable of
stimulating allogeneic MLR) characteristics similar to those of
TNF--induced mDCs (data not shown). As shown by Fig. 5A, mDCs induced by
anti-CD40 Ab did not migrate to fMLP (•), yet they still migrated
in response to C5a (
). In agreement with the chemotaxis data (Fig. 5A), iDCs expressed both FPR (Fig. 5B, lane
1) and C5aR (Fig. 5C, lane 1) mRNAs. Upon
anti-CD40 Ab-induced maturation, mDCs down-regulated FPR mRNA (Fig. 5B, lane 2) while maintaining a comparable level
of C5aR mRNA expression (Fig. 5C, lane 2).
Therefore, DC maturation induced by either TNF- or CD40 ligation
results in selective down-regulation of FPR, but not C5aR,
expression.
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Human DCs can be generated in vitro in a large number from either
monocytes or CD34+ HPC (11, 12, 13, 14, 15, 21, 22, 24, 25, 26, 27, 28, 33, 34). To address whether HPC-derived DCs behave
similarly to monocyte-derived DCs, we investigated the response to fMLP
and C5a as well as FPR and C5aR expression of HPC-derived DCs before
and after maturation. Human HPC-derived iDCs were
CD1alow, CD83-,
CD86low, and HLA-DRmedium
(Table III) and lacked the capacity to
stimulate allogeneic MLR (Table IV),
whereas HPC-derived mDCs were CD1ahigh,
CD83+, CD86high, and
HLA-DRhigh (Table III) with potent capacity to
stimulate allogeneic MLR (Table IV), indicating that they exhibited
immature and mature phenotypes, respectively. Interestingly, human
HPC-derived iDCs migrated, similar to monocyte-derived iDCs, toward
both fMLP (Fig. 6A, 
) and
C5a (Fig. 6A, 
). After TNF--induced maturation,
CD34+ HPC-derived mDCs migrated toward C5a (Fig. 6A, ), but not to fMLP (Fig. 6A, •).
Furthermore, CD34+ HPC-derived iDCs mobilized
intracellular Ca2+ in response to both fMLP and
C5a, whereas mDCs did so only in response to C5a, but not to fMLP (Fig. 6B). Concomitantly, HPC-derived iDCs expressed both FPR
(Fig. 6C, left, lane 1) and C5aR (Fig. 6C, right, lane 1) mRNAs, whereas mDCs
maintained comparable levels of C5aR mRNA (Fig. 6C,
right, lane 2), but down-regulated FPR mRNA to an
undetectable level (Fig. 6C, left, lane
2). Thus, upon maturation, human HPC-derived DCs also
differentially regulate their responsiveness to and receptor expression
for fMLP and C5a.
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It has been reported that induction of iDC migration by fMLP and
C5a is based on chemotaxis (5, 10, 36). To assure whether
C5a-induced mDC migration is due to chemotaxis or chemokinesis,
checkerboard analysis was performed. As shown by Table V, addition of C5a into the upper wells
alone did not cause mDCs to migrate across the membrane (row
1), indicating that C5a did not increase chemokinesis of mDCs.
Addition of C5a into the lower wells alone resulted in dose-dependent
mDC migration (column 1). Moreover, when C5a was added into
both the upper and lower wells, mDC migration was inhibited to various
degrees, depending on C5a concentrations in the lower and upper wells
(desensitization). Taken together, checkerboard analysis indicates that
C5a-induced mDC migration is also based on chemotaxis.
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Murine iDCs express CXCR4 (49), CCR1 (49, 50), CCR2 (49), and CCR5 (34, 49)
whereas murine mDCs are CXCR4-positive (49) and
CCR7-positive (49, 50, 51, 52). Unlike human iDCs, murine iDCs do
not express CCR4 (49) and CCR6 (50),
highlighting several differences between human and mouse DCs. This led
us to investigate whether murine DCs respond similarly to maturation
signals as do human DCs with particular regard to the regulation of
responsiveness to fMLP and C5a. As demonstrated by Fig. 7, C5a induced the migration of both
murine iDCs () and mDCs (). However, fMLP was only chemotactic
for murine iDCs (), but not for murine mDCs (•), suggesting that
maturation of murine DCs also differentially regulates their
responsiveness to fMLP and C5a. The optimal chemotactic concentration
of fMLP for murine iDCs was 10-6 M, which was
100-fold higher than that for human iDCs (10-8 M). This
difference presumably reflects the different sensitivities of human and
murine FPR to fMLP rather than the difference between human and murine
iDCs because 1) to induce comparable level of intracellular
Ca2+ mobilization in murine neutrophils, 100-fold or more
fMLP is needed than with human neutrophils (53); 2) about
100-fold more fMLP is required to induce comparable level of
intracellular Ca2+ mobilization in murine as compared with
human FPR-expressing Xenopus oocytes (53); and
3) human embryonic kidney 293 cells expressing murine FPR migrate in
response to fMLP with an optimal concentration of 10-6 M
(54), whereas those expressing human FPR migrate in
response to fMLP with an optimal concentration of 10-8 M
(55).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Upon maturation induced by TNF- or CD40 ligation, mDCs
down-regulated their responsiveness to and receptor expression for fMLP
while maintaining their responsiveness to and receptor expression for
C5a. These results are compatible with the finding that human
skin-derived DCs up-regulate their responsiveness to C5a after
treatment with TNF- for 24 h (36). Moreover, the
differential regulation of mDCs responsiveness to fMLP and C5a
paralleled the expression of FPR and C5aR. This differential regulation
seems to be DC maturation-dependent rather than TNF--dependent
because 1) DCs matured by CD40 ligation also exhibited the same
pattern, and 2) TNF- has been shown to promote, rather than
suppress, FPR expression in human neutrophils (59).
Because the expression of FPR mRNA was also down-regulated, it can be
speculated that a reduction in either FPR mRNA stability and/or FPR
gene transcription is responsible. FPR and C5aR genes are known to
cluster at the same narrow region of a chromosome (19 in human and 17
in mouse) (60, 61, 62), however, the structure of their
promoter regions is not well understood. Furthermore, how the
expression of FPR and C5aR genes is controlled remains unknown.
Therefore, how and why DC maturation results in differential regulation
of FPR and C5aR expression needs further investigation.
DCs generated from CD34+ HPC are heterogeneous. At least two subsets of DCs, CD1a+ and CD1a-, can be derived from CD34+ HPC-derived DCs (63). Although our results show that, similar to monocyte-derived DCs, human HPC-derived DCs also differentially down-regulate FPR, but not C5aR, expression upon maturation, whether different subsets of HPC-derived DCs behave similarly or differently in the course of maturation in terms of FPR and C5aR expression awaits further investigation.
Interaction of two chemokine receptors, CXCR4 and CCR7 that are known
to be expressed by mDCs (14, 21, 23, 33, 34, 35), with their
ligands, e.g., SDF-1, secondary lymphoid chemokine (also known as
6Ckine, Exodus-2, or TCA4), and EBI1 ligand chemokine (also known as
macrophage inflammatory protein-3ß), is involved in the recruitment
of mDCs to lymphoid tissues (35, 51, 64). This leads us to
propose that the interaction of C5a and C5aR may also participate in
the recruitment of mDCs to lymphoid tissues, specifically in guiding
and/or sorting mDCs to B cell follicles, where naive B cells acquire
native Ags delivered by mDCs (57, 65). Besides
macrophages, B cells have recently been found to be a source of C5a in
secondary lymphoid tissue (66). Follicular DCs, a
particular type of DC localizing in B cell follicles, retain Ag-Ab
complexes on their surfaces (67). Therefore, C5a may be
generated via the classical pathway of complement activation triggered
by follicular DC-bound immune complexes, thereby forming a C5a
gradient. C5a is also a chemoattractant for B cells (66, 68). Thus, locally generated C5a gradient may attract both mDCs
and naive B cells to B cell follicles to facilitate Ag transfer
(56, 57, 65).
Collectively, our results suggest that the interaction of FPR with its ligands is possibly involved in the recruitment of iDCs whereas that of C5a and C5aR may participate in the recruitment of both iDCs and mDCs in vivo to distinct anatomical sites. The observation that murine DC responded to fMLP and C5a similarly to human DCs indicates that mouse models can be used to decipher the in vivo roles of fMLP and C5a in DC trafficking, particularly by the use of recently established FPR and C5aR knockout mice (69, 70).
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Acknowledgments |
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Footnotes |
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2 The content of this publication does not necessarily reflect the views or policies of the Department of Health and Human Services, nor does mention of trade names, commercial products, or organizations imply endorsement by the U.S. Government. The publisher or recipient acknowledges right of the U.S. Government to retain a nonexclusive, royalty-free license in and to any copyright covering the article.
3 D.Y. and Q.C. contributed equally to this study.
4 Address correspondence and reprint requests to Dr. Joost J. Oppenheim, Laboratory of Molecular Immunoregulation, Division of Basic Sciences, National Cancer Institute-Frederick Cancer Research and Development Center, Building 560, Room 21-89, Frederick, MD 21702-1201.
5 Abbreviations used in this paper: DCs, dendritic cells; mDC, mature DC; iDC, immature DC; C5aR, C5a receptor; FPR, formyl peptide receptor; HPC, hemopoietic progenitor cells; rh, recombinant human; SDF-1, stromal cell-derived factor 1.
6 O. M. Z. Howard, H. F. Dong, J. Subleski, S. Strobl, A.-K. Shirakawa, J. J. Oppenheim, and E. L. Nelson. TARC and I-309 utilize CCR8 to induce chemotaxis of a CD83- subset of human monocyte-derived dendritic cells. Submitted for publication.
Received for publication February 4, 2000. Accepted for publication June 12, 2000.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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from lung dendritic cells. J. Exp. Med. 186:825. and is highly expressed in human dendritic cells. J. Exp. Med. 186:837. and IL-4. J. Leukocyte Biol. 66:837.[Abstract]
: modification of the current "maturation" paradigm and implication of in vivo immunoregulation. FASEB J. 13:2021. regulates expression of receptors for formyl-methionyl-leucyl-phenylalanine, leukotriene B4, and platelet-activating factor: dissociation from priming in human polymorphonuclear neutrophils. J. Immunol. 147:3842.[Abstract]
. J. Exp. Med. 184:695.This article has been cited by other articles:
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  Q. Peng, K. Li, K. Anderson, C. A. Farrar, B. Lu, R. A. G. Smith, S. H. Sacks, and W. Zhou Local production and activation of complement up-regulates the allostimulatory function of dendritic cells through C3a-C3aR interaction Blood, February 15, 2008; 111(4): 2452 - 2461. [Abstract] [Full Text] [PDF]
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 D. Yang, Q. Chen, S. B. Su, P. Zhang, K. Kurosaka, R. R. Caspi, S. M. Michalek, H. F. Rosenberg, N. Zhang, and J. J. Oppenheim Eosinophil-derived neurotoxin acts as an alarmin to activate the TLR2-MyD88 signal pathway in dendritic cells and enhances Th2 immune responses J. Exp. Med., January 21, 2008; 205(1): 79 - 90. [Abstract] [Full Text] [PDF]
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 A. Karlsson, E. Nygren, J. Karlsson, I. Nordstrom, C. Dahlgren, and K. Eriksson Ability of Monocyte-Derived Dendritic Cells To Secrete Oxygen Radicals in Response to Formyl Peptide Receptor Family Agonists Compared to That of Myeloid and Plasmacytoid Dendritic Cells Clin. Vaccine Immunol., March 1, 2007; 14(3): 328 - 330. [Abstract] [Full Text] [PDF]
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 M. A. Connelly, R. A. Moulton, A. K. Smith, D. R. Lindsey, M. Sinha, R. A. Wetsel, and C. Jagannath Mycobacteria-primed macrophages and dendritic cells induce an up-regulation of complement C5a anaphylatoxin receptor (CD88) in CD3+ murine T cells J. Leukoc. Biol., January 1, 2007; 81(1): 212 - 220. [Abstract] [Full Text] [PDF]
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 D. J. Klinke II An age-structured model of dendritic cell trafficking in the lung. Am J Physiol Lung Cell Mol Physiol, November 1, 2006; 291(5): L1038 - L1049. [Abstract] [Full Text] [PDF]
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H. L. Rittner, S. A. Mousa, D. Labuz, K. Beschmann, M. Schafer, C. Stein, and A. Brack Selective local PMN recruitment by CXCL1 or CXCL2/3 injection does not cause inflammatory pain J. Leukoc. Biol., May 1, 2006; 79(5): 1022 - 1032. [Abstract] [Full Text] [PDF]
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D. F. Legler, P. Krause, E. Scandella, E. Singer, and M. Groettrup Prostaglandin E2 Is Generally Required for Human Dendritic Cell Migration and Exerts Its Effect via EP2 and EP4 Receptors J. Immunol., January 15, 2006; 176(2): 966 - 973. [Abstract] [Full Text] [PDF]
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K. Kurosaka, Q. Chen, F. Yarovinsky, J. J. Oppenheim, and D. Yang Mouse Cathelin-Related Antimicrobial Peptide Chemoattracts Leukocytes Using Formyl Peptide Receptor-Like 1/Mouse Formyl Peptide Receptor-Like 2 as the Receptor and Acts as an Immune Adjuvant J. Immunol., May 15, 2005; 174(10): 6257 - 6265. [Abstract] [Full Text] [PDF]
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D. Yang, Q. Chen, H. F. Rosenberg, S. M. Rybak, D. L. Newton, Z. Y. Wang, Q. Fu, V. T. Tchernev, M. Wang, B. Schweitzer, et al. Human Ribonuclease A Superfamily Members, Eosinophil-Derived Neurotoxin and Pancreatic Ribonuclease, Induce Dendritic Cell Maturation and Activation J. Immunol., November 15, 2004; 173(10): 6134 - 6142. [Abstract] [Full Text] [PDF]
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A. Biragyn, P. A. Ruffini, M. Coscia, L. K. Harvey, S. S. Neelapu, S. Baskar, J.-M. Wang, and L. W. Kwak Chemokine receptor-mediated delivery directs self-tumor antigen efficiently into the class II processing pathway in vitro and induces protective immunity in vivo Blood, October 1, 2004; 104(7): 1961 - 1969. [Abstract] [Full Text] [PDF]
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A. H. J. Kim, I. D. Dimitriou, M. C. H. Holland, D. Mastellos, Y. M. Mueller, J. D. Altman, J. D. Lambris, and P. D. Katsikis Complement C5a Receptor Is Essential for the Optimal Generation of Antiviral CD8+ T Cell Responses J. Immunol., August 15, 2004; 173(4): 2524 - 2529. [Abstract] [Full Text] [PDF]
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Q. Chen, D. Wade, K. Kurosaka, Z. Y. Wang, J. J. Oppenheim, and D. Yang Temporin A and Related Frog Antimicrobial Peptides Use Formyl Peptide Receptor-Like 1 as a Receptor to Chemoattract Phagocytes J. Immunol., August 15, 2004; 173(4): 2652 - 2659. [Abstract] [Full Text] [PDF]
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A. J. Melendez and F. B. M. Ibrahim Antisense Knockdown of Sphingosine Kinase 1 in Human Macrophages Inhibits C5a Receptor-Dependent Signal Transduction, Ca2+ Signals, Enzyme Release, Cytokine Production, and Chemotaxis J. Immunol., August 1, 2004; 173(3): 1596 - 1603. [Abstract] [Full Text] [PDF]
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D. Yang, H. F. Rosenberg, Q. Chen, K. D. Dyer, K. Kurosaka, and J. J. Oppenheim Eosinophil-derived neurotoxin (EDN), an antimicrobial protein with chemotactic activities for dendritic cells Blood, November 1, 2003; 102(9): 3396 - 3403. [Abstract] [Full Text] [PDF]
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A. Soruri, Z. Kiafard, C. Dettmer, J. Riggert, J. Kohl, and J. Zwirner IL-4 Down-Regulates Anaphylatoxin Receptors in Monocytes and Dendritic Cells and Impairs Anaphylatoxin-Induced Migration In Vivo J. Immunol., March 15, 2003; 170(6): 3306 - 3314. [Abstract] [Full Text] [PDF]
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M. Suresh, H. Molina, M. S. Salvato, D. Mastellos, J. D. Lambris, and M. Sandor Complement Component 3 Is Required for Optimal Expansion of CD8 T Cells During a Systemic Viral Infection J. Immunol., January 15, 2003; 170(2): 788 - 794. [Abstract] [Full Text] [PDF]
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D. Yang, Q. Chen, B. Gertz, R. He, M. Phulsuksombati, R. D. Ye, and J. J. Oppenheim Human dendritic cells express functional formyl peptide receptor-like-2 (FPRL2) throughout maturation J. Leukoc. Biol., September 1, 2002; 72(3): 598 - 607. [Abstract] [Full Text] [PDF]
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M. Idzko, S. Dichmann, D. Ferrari, F. Di Virgilio, A. la Sala, G. Girolomoni, E. Panther, and J. Norgauer Nucleotides induce chemotaxis and actin polymerization in immature but not mature human dendritic cells via activation of pertussis toxin-sensitive P2y receptors Blood, July 18, 2002; 100(3): 925 - 932. [Abstract] [Full Text] [PDF]
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 S. SAADI, L. E. WRENSHALL, and J. L. PLATT Regional manifestations and control of the immune system FASEB J, June 1, 2002; 16(8): 849 - 856. [Abstract] [Full Text] [PDF]
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 M. F. Lipscomb and B. J. Masten Dendritic Cells: Immune Regulators in Health and Disease Physiol Rev, January 1, 2002; 82(1): 97 - 130. [Abstract] [Full Text] [PDF]
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A. Biragyn, M. Surenhu, D. Yang, P. A. Ruffini, B. A. Haines, E. Klyushnenkova, J. J. Oppenheim, and L. W. Kwak Mediators of Innate Immunity That Target Immature, But Not Mature, Dendritic Cells Induce Antitumor Immunity When Genetically Fused with Nonimmunogenic Tumor Antigens J. Immunol., December 1, 2001; 167(11): 6644 - 6653. [Abstract] [Full Text] [PDF]
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D. Yang, O. Chertov, and J. J. Oppenheim Participation of mammalian defensins and cathelicidins in anti-microbial immunity: receptors and activities of human defensins and cathelicidin (LL-37) J. Leukoc. Biol., May 1, 2001; 69(5): 691 - 697. [Abstract] [Full Text]
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D. Yang, Q. Chen, Y. Le, J. M. Wang, and J. J. Oppenheim Differential Regulation of Formyl Peptide Receptor-Like 1 Expression During the Differentiation of Monocytes to Dendritic Cells and Macrophages J. Immunol., March 15, 2001; 166(6): 4092 - 4098. [Abstract] [Full Text] [PDF]
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De Yang, Q. Chen, A. P. Schmidt, G. M. Anderson, J. M. Wang, J. Wooters, J. J. Oppenheim, and O. Chertov Ll-37, the Neutrophil Granule-And Epithelial Cell-Derived Cathelicidin, Utilizes Formyl Peptide Receptor-Like 1 (Fprl1) as a Receptor to Chemoattract Human Peripheral Blood Neutrophils, Monocytes, and T Cells J. Exp. Med., October 2, 2000; 192(7): 1069 - 1074. [Abstract] [Full Text] [PDF]
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