* Laboratory of Molecular Immunoregulation, Center for Cancer Research, National Cancer Institute at Frederick, National Institutes of Health, Maryland; and
Department of Pharmacology, College of Medicine, University of Illinois, Chicago
Correspondence: Dr. Joost J. Oppenheim, LMI, CCR, NCI at Frederick, Building 560, Room 21-89, Frederick, MD 21702-1201. E-mail: oppenhei{at}mail.ncifcrf.gov
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: chemotaxis • Ca2+ mobilization • maturation
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Regulation of DC trafficking in vivo like trafficking of other leukocytes is influenced by many chemotactic factors and adhesion molecules [2 , 4 ]. However, the capacity of iDC and mDC to migrate to different anatomical sites is primarily determined by their differential expression of G-protein-coupled seven-transmembrane domain receptors (GPCRs) specific for chemokines and classical chemoattractants [1 , 2 , 4 , 5 ]. Of the 20 chemokine receptors identified so far, in vitro studies have shown that iDC express CXC chemokine receptor (CXCR)1, 2, 4 and CCR1–6 and are able to migrate in response to their respective ligands [5 6 7 ]. In contrast, mDC express only CXCR4 and CCR7 and thus are able to migrate, respectively, in response to stromal cell-derived factor-1/CXCL12 and CCR7 ligands including 6Ckine/secondary lymphoid chemokine/exodus-2/CCL21 and macrophage inflammatory protein-3ß/EBI1 ligand chemokine/exodus-3/CCL19 [2 , 5 , 8 ]. The in vivo contribution of CCR6 to iDC migration and that of CCR7 to mDC trafficking has been clearly demonstrated [9 10 11 ], indicating that in vitro investigation of the expression of chemotactic receptors by DC provides a useful step for defining their involvement in DC trafficking.
The receptors for classical chemoattractants may also regulate the trafficking of DC precursors and DC. Classical chemoattractant receptors include C3a and C5a receptors, platelet-activating factor receptor (PAFR), and the receptors for formyl peptides (FPR). Human monocyte-derived myeloid iDC and mDC express functional PAFR [12 ]. Human DC derived from monocytes or CD34+ progenitors express functional C5a receptor even after maturation [13 ], indicating the interaction of C5a and its receptor may regulate trafficking of iDC and mDC. Indeed, it has been demonstrated that C5a is required for the induction of contact sensitivity in mice [14 15 16 ]. The human FPR subfamily has three members including FPR and two additional homologues termed FPR-like 1 and 2 (FPRL1 and FPRL2) [17 ]. We have previously shown that human iDC express functional FPR, and its expression is down-regulated on mDC [13 ]. We have recently demonstrated that FPRL1 is down-regulated even as DC precursors differentiate into iDC [18 ]. Whether human DC express FPRL2 has not been reported.
Human FPR was cloned in 1990 [19 ] and uses the prototype formyl peptide N-formyl-Met-Leu-Phe (fMLP) as a high affinity agonist [17 , 20 21 22 ]. Human FPRL1 (also known as FPR2 or FPRH1) was cloned in 1992 by several independent groups [23 24 25 ]. Although sharing 69% amino acid identity to FPR, FPRL1 only has very low affinity for fMLP [17 , 24 ]. Subsequently, FPRL1 was reported to be a functional, high affinity receptor for lipoxin A4 and was given another name, LXA4R [26 ]. In recent years, a number of diverse peptides/proteins including MMK-1, a peptide isolated from a random peptide library [27 ], have been reported to act as FPRL1-selective agonistic ligands [28 29 30 31 ]. The hexapeptide Trp-Lys-Tyr-Met-Val-D-Met (WKYMVm), originally identified from a combinatorial peptide library by its capability to stimulate phosphoinositide hydrolysis in lymphocyte cell lines [32 ], is a potent leukocyte activator [33 34 35 ] and has been shown to act on human FPR and FPRL1 [36 , 37 ]. Human FPRL2, also cloned in 1992, shares 56% and 83% amino acid identity with FPR and FPRL1, respectively [23 , 25 ]. FPRL2 mRNA is expressed by monocytes, but not by neutrophils [38 ]. Cells transfected to overexpress FPRL2 have recently been shown to respond to WKYMVm [39 ]. However, it is not clear whether functional FPRL2 is expressed by primary cells.
In the course of investigating the regulation of FPR and FPRL1 during human myeloid DC differentiation and maturation [13 , 18 ], we observed that mDC, although unable to respond to the fMLP, could nevertheless migrate in response to WKYMVm. As mDC exhibited no functional FPR and FPRL1 expression [13 , 18 ], we speculated that mDC must express another receptor that enabled them to respond to WKYMVm. Searching for this receptor, we found that human myeloid DC express FPRL2 at mRNA and protein levels and maintain FPRL2 expression even after maturation. We also demonstrated that FPRL2 expressed by DC was functional and mediated the effect of WKYMVm on mDC. Thus, we propose that the interaction of FPRL2 with its unknown endogenous or exogenous ligand(s) may be involved in regulating trafficking of iDC to sites of Ag deposition and of mDC from peripheral tissues to secondary lymphoid organs.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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(rhTNF-; sp.
Act.=2x107 U/mg), rh granulocyte macrophage-colony
stimulating factor (GM-CSF; sp. Act.=107 U/mg), and rh
interleukin (IL)-4 (sp. Act.=2x106 U/mg) were purchased
from PeproTech (Rocky Hill, NJ). Fluorescein isothiocyanate
(FITC)-conjugated goat anti-rabbit immunoglobulin G (IgG) antibody,
synthetic fMLP, Escherichia coli lipopolysaccharide (LPS;
serotype 026:B5), and pertussis toxin (PTX) were purchased from Sigma
Chemical Co. (St. Louis, MO). The other antibodies used for flow
cytometry were purchased from BD PharMingen (San Diego, CA). Polyclonal
anti-FPRL1 and 2 antiserum was raised in a rabbit against a synthetic
peptide AANSASPPAETELQAM, which corresponds to the carboxyl terminal 16
amino acid residues of FPRL1 [23
, 24
]. As a
result of the high homology of the carboxyl terminus of FPRL1 with that
of FPRL2 (12 out of the last 16 and 9 out of the last 10 amino acids
are identical) [17
, 23
24
25
], the antiserum
generated was found to cross-react with the corresponding carboxyl
terminus of FPRL2 by flow cytometry analysis (R. He and R. D. Ye,
unpublished results). MMK-1, an FPRL1-specific ligand of 13 amino acid
(LESIFRSLLFRVM) [27
], and the hexapeptide WKYMVm (m
represents a D-Met residue) [32
] were
synthesized and purified by the Department of Biochemistry, Colorado
State University (Fort Collins). The purity for both peptides was more
than 90%, and the amino acid composition was verified by mass
spectrometry.
Isolation of human peripheral blood monocytes and DC culture
Human peripheral blood mononuclear cells (PBMC) were isolated
from leukopacks (Courtesy of the Transfusion Medicine Department, NIH
Clinic Center, Bethesda, MD) by routine Ficoll-Hypaque density gradient
centrifugation. Monocytes were purified from human PBMC with the use of
the magnetic cell sorter CD14 monocyte isolation kit (Miltenyi Biotech
Inc., Auburn, CA), according to the manufacturer’s recommendation. The
purity of monocytes was checked by FACScan analysis. Monocyte
preparation with purity less than 95% was not used. The generation of
DC from monocytes was carried out as described previously
[13
, 18
]. Briefly, monocytes were
differentiated to iDC by incubating 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 (50 ng/ml) at 37°C in a humidified
CO2 (5%) incubator for 7 days. To induce DC maturation,
iDC were cultured in the same cytokine cocktails with added rhTNF-
(50 ng/ml) or LPS (100 ng/ml) for 48 h at 37°C in a humidified
CO2 (5%) incubator. As confirmed by flow cytometry
analysis, iDC were CD1a+, CD14-,
CD40low, CD83-, CD86low, human
leukocyte Ag (HLA)-DRmedium, whereas mDC were
CD1a+, CD14-, CD40high,
CD83+, CD86high, and HLA-DRhigh
(data not shown; and refs. [13
, 18
]).
Chemotaxis assay
Migration of monocytes and DC in response to chemotactic factors
was assessed using a 48-well microchemotaxis chamber technique as
previously described [6
, 13
]. Briefly,
different concentrations of chemotactic factors were placed in
triplicates in the wells of the lower compartment of the chamber (Neuro
Probe, Cabin John, MD), and cells (106 cells/ml) were added
to wells of the upper compartment. In some experiments, various
concentrations of chemotactic factor were also included in the wells of
the upper compartment. The lower compartment was separated from the
upper compartment 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
chemotactic index (C.I.), which is defined as the "fold" increase
in the number of migrating cells in the presence of test factors over
the spontaneous cell migration (in the absence of test factors). The
statistical significance of the increase in cell migration was
determined by an unpaired t-test.
Measurement of calcium flux
Monocytes and DC (107 cells/ml in RPMI 1640
containing 10% FBS) were loaded by incubating with 5 µM Fura-2
(Molecular Probes, Eugene, OR) at 24°C for 30 min in the dark.
Subsequently, the cells were washed with 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% bovine serum
albumin (BSA), pH 7.4]. Each 2 ml cell suspension was then transferred
into a quartz cuvette, which was placed in a luminescence spectrometer
LS50 B (Perkin-Elmer Limited, Beaconsfield, UK). 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. In some experiments, loaded
cells were treated with PTX (final concentration of 200 ng/ml) at
37°C for 30 min and were washed twice with saline buffer before
measuring the calcium flux of the cells in response to chemotactic
factors.
Flow cytometry
DC (106/sample) were washed three times with
phosphate-buffered saline (PBS) and fixed for 10 min at room
temperature in 3.7% paraformaldehyde (W/V in PBS). Subsequently, the
cells were permeabilized/blocked by treatment with
permeabilizing/blocking (P/B) buffer (PBS containing 0.1% saponin, 1%
BSA, and 10 µg/ml human IgG). Thereafter, the cells were stained with
rabbit preimmune serum or rabbit anti-FPRL1 and 2 immune serum (both at
1:400 dilution with P/B buffer) at room temperature for 1 h. After
washing three times with P/B buffer, the cells were further stained
with FITC-conjugated goat anti-rabbit IgG (1:40 dilution with P/B
buffer) for 30 min at room temperature. The stained cells were washed
twice with P/B buffer, twice with PBS, fixed with 1% paraformaldehyde
in PBS at 4°C overnight, and analyzed the next day with a flow
cytometer (Coulter Epics®, Miami, FL).
RNA isolation and reverse transcriptase-polymerase chain reaction
(RT-PCR)
Total RNA from monocytes, DC, and macrophages was isolated by
the use of TRIzol® Reagent (Life Technologies, Grand Island, NY). The
RNAs were cleaned by treatment with RNase-free DNase I (Stratagene, La
Jolla, CA). RT-PCR was performed by the use of the ProSTARTM HF
single-tube RT-PCR system (Stratagene). Briefly, 5–50 ng total RNA was
used in the RT-PCR reaction. After reverse transcription at 37°C for
15 min and inactivation of Moloney murine leukemia virus reverse
transcriptase at 95°C for 1 min, FPR, FPRL2, and glyceraldehyde
3-phosphate dehydrogenase (GAPDH) cDNA fragments were amplified by 40
cycles of PCR (denature at 95°C for 30 s, annealing at 60°C
for 30 s, and extension at 68°C for 2 min) reaction, and the
last extension was performed at 68°C for 10 min. The sense and
antisense primers for FPR were 5'-CTCCAGTTGGACTAGCCACA-3' (nucleotides
1639
1658 in the coding region of exon 2) and
5'-CCATCACCCAGGGCCCAATG-3' (nucleotides 53415359 in the coding
region of exon 3), respectively, which resulted in the amplification of
a 500-bp FPR-specific cDNA fragment as previously reported
[40
]. The sense and antisense primers for FPRL2 were
5'-GCCAAGGTCTTTCTGATCC-3' (nucleotides 592610 as counted from the
starting position of the open reading frame) and
5'-GGTCTGGGCTGAGTCAGGGA-3' (nucleotides 977996), which enabled the
amplification of a 404-bp FPRL2-specific cDNA fragment as verified by
sequencing. The primers for FPRL1 were 5'-CTGCTGGTGCTGCTGGCAAG-3' and
5'-AATATCCCTGACCCCATCCTCA-3', which, as previously shown, enabled the
amplification of a 1.1-kb FPRL1-specific cDNA fragment
[18
]. The primers for human GAPDH were
5'-GATGACATCAAGAAGGTGGTGAA-3' and 5'-GTCTTACTCCTTGGAGGCCATGT-3', which
resulted in the amplification of a fragment of 246 bp as previously
described [13
, 18
]. PCR products were
identified on 2% agarose gel, stained with ethidium bromide, and
photodocumented after washing off excessive dye with water.
Confocal microscopy
iDC and mDC were seeded into gelatin-coated, glass-bottom, 35-mm
dishes (MatTek Corp., Ahland, MA; Cat. No. P35GC-0-14-C) and were
incubated in appropriate medium at 37°C for 24 h prior to the
experiments. Cycloheximide was added to a final concentration of 100
µg/ml during the last 3 h of incubation to stop protein
synthesis. Twenty minutes before the end of the incubation period,
WKYMVm was added to some dishes to a final concentration of 10 µM to
induce receptor internalization. The cells were then rinsed three times
with ice-cold PBS to remove ligand and FCS and were fixed in 3.7%
paraformaldehyde at room temperature for 10 min. After three rinses
with PBS, the cells were permeabilized with PBS containing 0.1%
saponin and 0.02% Nonidet P-40 at room temperature for 30 min. The
permeabilized cells were sequentially stained with the primary
polyclonal antibody (rabbit preimmune serum or anti-FPRL1 and 2; both
were diluted by 1:500 in PBS containing 0.1% saponin and 10 µg/ml
human IgG) and the secondary FITC-labeled goat anti-rabbit IgG (at
1:100 dilution). The stained cells were extensively washed, air-dried,
and covered with several drops of Gel/MountTM (Biomeda Corp., Foster
City, CA). A confocal laser-activated scanning microscope (Carl Zeiss
410) was used for the microscopic analysis of stained cells.
FITC-stained cells were excited at 488 nm, and the fluorescence emitted
between 510 nm and 565 nm was collected.
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and examined. Interestingly, mDC,
although unable to migrate to fMLP or MMK-1, nevertheless migrated in
response to WKYMVm (Fig. 1
, upper right panel). Almost identical
results were obtained when mDC generated in the presence of LPS instead
of TNF- were used for the chemotaxis (Fig. 1
, lower right panel).
Thus, mDC generated in the presence of TNF- were used in subsequent
experiments.
 View larger version (30K): [in a new window] |
Figure 1. Migration of monocytes and monocyte-derived DC in response to fMLP,
MMK-1, and WKYMVm. Human peripheral blood monocytes were isolated from
a single donor and incubated in the presence of rhGM-CSF and rhIL-4 for
7 days in humidified air containing 5% CO2 to generate
iDC. mDC were generated by culturing iDC in the presence of rhGM-CSF,
rhIL-4, and rhTNF- or LPS for an additional 2 days. The migration of
monocytes, iDC, and mDC in response to various concentrations of fMLP,
MMK-1, and WKYMVm was tested by the use of 48-well chemotaxis chambers,
and the results are shown as the average C. I.
(mean±SD) of triplicated wells. The spontaneous migration
(in the absence of chemotactic factors) for monocytes and DC was
3050 cells and 5070 cells per high-powered field, respectively.
Similar results were obtained using cells of more than five individual
donors.
|
 View larger version (50K): [in a new window] |
Figure 2. Checkerboard analysis of WKYMVm-induced mDC migration. Various
concentrations of WKYMVm were added to the lower wells, and mDC in the
absence or presence of various concentrations of WKYMVm, were added to
the upper wells of a chamber. After a 90-min incubation at 37°C, the
membrane was removed, scraped, stained, and dried. The migration of mDC
to the lower surface of the membrane was determined and shown as the
average C. I. (mean±SD) of triplicated wells. Two
separate experiments showed almost identical results.
|
 View larger version (24K): [in a new window] |
Figure 3. Ca2+ mobilization of monocytes and monocyte-derived DC in
response to fMLP, MMK-1, and WKYMVm. Monocytes and monocyte-derived iDC
and mDC were from the same donors. Stimulants (fMLP, MMK-1, and WKYMVm)
were added 36 s after starting the recording at the final
concentration (M) as specified. One representative experiment out of
three is shown.
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 View larger version (21K): [in a new window] |
Figure 4. Inhibition by PTX of the effect of WKYMVm on mDC. (A) Chemotaxis. mDC
were treated in the absence (dotted bar) or presence (hatched bar) of
PTX at 200 ng/ml at 37°C for 30 min before performing chemotaxis
assay. *, P < 0.05; **, P < 0.01. (B)
Ca2+ flux. Fura-2-loaded mDC were incubated with or without
200 ng/ml PTX (PTX+ and PTX-, respectively), washed once with saline
buffer, and examined for their Ca2+ mobilization in
response to WKYMVm. Arrows indicate the time points where WKYMVm were
added at the final concentrations as specified. PTX treatment of DC did
not decrease their viability (data not shown) or motility, as the
random migration of treated mDC was similar to untreated cells [the
first two bars (A)]. Two separate experiments showed almost identical
results.
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 View larger version (56K): [in a new window] |
Figure 5. Expression of FPRL2 mRNA by iDC and mDC. Total RNA was isolated
from iDC and mDC derived from the same donor. (A) Total iDC or mDC RNAs
(50 or 5 ng) were used for the RT-PCR amplification. RT-PCR products of
FPR (top), FPRL2 (middle), and GAPDH (bottom) were displayed by agarose
gel (2%) electrophoresis. The marker used was a 50-bp DNA ladder. (B)
Each 5 ng total RNA derived from monocytes (used as a positive
control), iDC, and mDC was used for the amplification of FPRL1 and
GAPDH by RT-PCR. The products of FPRL1 (top) and GAPDH (bottom) were
displayed similarly as in Figure 1A
. The marker used was a 1-kb DNA
ladder. The anticipated size for FPR, FPRL1, FPRL2, and GAPDH was 500-,
1100-, 404-, and 246-bp, respectively. One representative experiment
out of three is shown.
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 View larger version (20K): [in a new window] |
Figure 6. FPRL2 protein expression by iDC and mDC. iDC and mDC were generated
from monocytes isolated from the same donor. Aliquots of
106 iDC or mDC were permeabilized and stained with rabbit
preimmune serum (open area) or anti-FPRL1 and 2 serum (solid area).
After subsequent staining with FITC-conjugated goat anti-rabbit IgG,
the cells were analyzed with a flow cytometer. Shown are the overlay
histograms for iDC (left panel) and mDC (right panel). Two separate
experiments showed almost identical results.
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FPRL2 expressed on DC is functional
To conclude that the FPRL2 expressed on mDC is the receptor that
mediates WKYMVm-induced mDC activation, direct evidence that FPRL2 on
mDC can transduce signal from WKYMVm is essential. We therefore
explored whether treatment of mDC with WKYMVm could induce the
internalization of FPRL2, one of the characteristic responses common to
all Gi protein-coupled chemotactic receptors upon agonist-induced
activation [42
]. To do so, iDC and mDC were generated
from the same population of precursors, treated at 37°C for 20 min in
the absence (sham) or presence of 10 µM WKYMVm, and immunostained for
confocal microscopic analysis. Although FPRL2-specific antibody has not
been available, we used the anti-FPRL1 and 2 antiserum to detect FPRL2
in DC, based on the fact that iDC and mDC do not express FPRL1 (Figs. 1 3
and 5
; and ref. [18
]). In iDC and mDC treated in
the absense of WKYMVm (sham), staining with anti-FPRL1 and 2 revealed
that FPRL2 was predominantly distributed at or close to the cell
surface, as illustrated by the bright fluorescent staining in the cell
periphery (Fig. 7
, left panels). The FPRL2 was unevenly distributed since a
punctated fluorescence at or underneath the cell membrane was observed.
In about 10% of the cells (iDC and mDC), fluorescent staining was
visualized within the cytoplasm (e.g., white arrow). Why FPRL2 is not
evenly distributed at the cell surface in DC is not clear, but a
similar pattern was also observed in FPRL2-transfected cells
[39
]. Importantly, treatment of iDC and mDC with WKYMVm
resulted in the accumulation of bright fluorescent staining
predominantly in the cytoplasm (red arrows), albeit the membrane of
some cells was still stained (Fig. 7
, right panels). iDC and mDC
stained with rabbit preimmune serum showed very faint green
fluorescence (data not shown), indicating that the bright fluorescence
observed in Figure 7
was specifically a result of anti-FPRL1
and 2 antibody rather than nonspecific binding of rabbit IgG in the
antiserum used. Thus, WKYMVm was able to trigger considerable FPRL2
internalization in iDC and mDC, suggesting that FPRL2 expressed by DC
is functional.
 View larger version (22K): [in a new window] |
Figure 7. WKYMVm-induced internalization of FPRL2 in iDC and mDC. iDC and mDC
generated from monocytes isolated from the same donor were treated
first at 37°C with cycloheximide (100 µg/ml) for 3 h and
subsequently at 37°C for 20 min in the absence or presence of 10 µM
WKYMVm (sham- and WKYMVm-treated, respectively). Thereafter, the cells
were fixed, permeabilized, and sequentially stained with primary
antibody (rabbit preimmune serum or anti-FPRL1 and 2 antiserum) and
FITC-conjugated secondary antibody for confocal microscopy. Shown are
the images of the medial plane of cells sham-treated or treated with
WKYMVm. The optical section thickness was calculated to be 0.69 µm.
DC stained with preimmune serum, as the primary antibody showed
extremely faint green fluorescence (data not shown). The white arrow in
the left lower panel indicates FPRL2 fluorescence inside a mDC. The red
arrows point to examples that FPRL2 fluorescence was accumulated inside
iDC and mDC. One representative experiment out of three is
shown.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Peripheral blood monocytes are the myeloid DC precursors that can be differentiated into DC in vitro, either by treatment with appropriate cytokines [43 , 44 ] or by incubation with endothelial cells grown on a collagen matrix [45 ], and in vivo [46 ]. Although it has not been directly demonstrated as a result of the unavailability of FPRL2-specific agonistic ligand or FPRL2-specific antibody, several lines of evidence indicate that monocytes, the precursors of myeloid DC, presumably also express functional FPRL2. First, monocytes express FPRL2 at mRNA level [38 ]. Secondly, FPRL2 expressed by monocyte-derived DC is functional. Finally, comparison of the Ca2+ mobilization signals induced by WKYMVm in monocytes, iDC, and mDC (Fig. 3) indicates that monocytes are most sensitive. This is presumably a result of the expression by monocytes of all three WKYMVm-responsive receptors, including FPR, FPRL1, and FPRL2. Thus, functional FPRL2 is expressed by human DC precursors (monocytes) and iDC, and its expression is maintained even after DC maturation.
Based on the results of our previous [13 , 18 ] and present studies, the functional expression of the three members of the human FPR subfamily can be summarized as in Table 1 . As the interaction of chemotactic receptor(s) on a particular leukocyte with corresponding ligand(s) regulates the trafficking of that cell [2 , 4 , 8 , 17 ], it can thus be hypothesized that all of the three receptors are differentially involved in the homing of DC precursors (monocytes), iDC, and mDC. Indeed, several previous studies suggest that the migration of monocytes to sites of inflammation and of iDC to sites of Ag deposition is regulated by FPR agonists [17 , 47 48 49 ]. The lack of FPRL1 expression by DC indicates that the interaction of FPRL1 with its endogenous ligands predominantly contributes to the recruitment of neutrophils, monocytes, and macrophages [18 , 36 , 38 ]. In addition to FPRL2, several other G-protein-coupled chemotactic receptors are expressed by mDC, including CCR7, CXCR4, C5a receptor, and PAFR [5 , 12 , 13 , 50 ]. The contribution of CCR7 and CXCR4 in regulating mDC trafficking has been clearly demonstrated [10 , 11 , 51 ]. C5a receptor may also contribute to mDC homing to regional lymph nodes, as mice lacking C5a or C5a receptor exhibit impaired contact sensitivity reaction [14 15 16 ]. Although mDC express only FPRL2 but not FPR or FPRL1, whether the interaction of FPRL2 on mDC with its endogenous ligand(s) may also contribute to regulating trafficking of mDC from peripheral tissues to secondary lymphoid organs remains to be investigated. The mouse as well as the human FPR subfamily is relatively well-characterized. In contrast to humans, the mouse FPR subfamily has six members [52 53 54 ]. Mouse FPRs share 77% amino acid identity with human FPRs and also use fMLP as an agonistic ligand [52 , 53 ]. Mouse FPRL1 is one ortholog for human FPRL1 that shares 74% amino acid identity and like human FPRL1, interacts with fMLP with low affinity [24 , 55 ] but acts as a high affinity receptor for LXA4 [26 , 56 ]. The gene product of Fpr-rs2 [52 ], recently named as mouse FPR2 [53 ], shares high homology with mouse FPRL1 (81% amino acid identity) and uses serum amyloid A (SAA) as an agonist [57 ]. Therefore, the functions of mouse FPRL1 and FPR2 are similar to that of human FPRL1. However, the mouse ortholog(s) for human FPRL2 has not been characterized.
|
View this table: [in a new window] |
Table 1. Functional Expression of FPR, FPRL1, and FPRL2 During the
Differentiation and Maturation of Human Myeloid DC
|
Thus, FPR may have evolved to sense prokaryotic microorganism-derived peptides (such as fMLP) to recruit FPR-expressing cells including iDC to sites of bacterial entry in the initial phase of infection and to receive annexin I signal to suppress the accumulation of FPR-expressing cells in the late phase of inflammation. Of about one dozen of the agonistic ligands identified so far for FPRL1 [18 , 24 , 26 27 28 29 30 31 , 36 , 37 , 63 64 65 66 ], six ligands are natural/endogenous, including LXA4 [26 , 63 ], SAA [28 ], LL-37, the cleaved carboxyl terminal domain of human antimicrobial protein cathelicidin/hCAP18 [31 ], the mitochondrial necrotactic peptide (MYFINILTL) derived from reduced nicotinamide adenine dinucleotide dehydrogenase subunit 1 [30 ], amyloid ß1-42 [65 ], and a cellular prion peptide fragment PrP106–126 [66 ]. These endogenous ligands are generated predominantly in association with inflammation and/or tissue injury [30 , 56 , 63 , 67 68 69 70 71 ]. The lack of FPRL1 expression by DC is noteworthy and may indicate that DC activation through FPRL1 in circumstances where endogenous FPRL1 ligands are generated could be potentially inimical to the host (e.g., by initiating unwanted autoimmune responses). Furthermore, the interaction of FPRL1 with its endogenous ligands may have a negative feedback-inhibitory effect on FPRL1-positive cells, especially when produced systemically in large amounts such as during severe infection. For example, LXA4 has been reported to act as a potent inhibitor of acute inflammation by suppressing CD11/18 expression and chemokine production of FPRL1-positive cells [56 , 63 , 72 ]. In addition, serum amyloid is able to inhibit the oxidative burst response of neutrophils as well as cell adhesion [73 , 74 ]. Scrapie prion protein has also been reported to inhibit neutrophil functions [75 ]. Thus, it would be counterproductive for DC to sense these ligands, as DC should not be inhibited even during severe systemic inflammation to perform Ag uptake, processing, and presentation. Although no endogenous and/or natural ligand(s) for FPRL2 has been identified as yet, they are presumably to be generated along the trafficking routes of DC precursors and DC.
In brief, we have demonstrated that FPRL2 is expressed by primary human iDC and mDC and suggest that the interaction of FPRL2 with its natural endogenous ligand(s) may participate in regulating DC trafficking. Direct investigation of the function of FPRL2, in particular its involvement in regulating DC trafficking, will become more feasible when the FPRL2-specific neutralizing antibody becomes available, the natural and/or endogenous agonist(s) for FPRL2 is identified, and the mouse ortholog(s) for human FPRL2 has been characterized.
Received January 7, 2002; revised March 22, 2002; accepted March 27, 2002.
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A. de Paulis, N. Prevete, I. Fiorentino, A. F. Walls, M. Curto, A. Petraroli, V. Castaldo, P. Ceppa, R. Fiocca, and G. Marone Basophils Infiltrate Human Gastric Mucosa at Sites of Helicobacter pylori Infection, and Exhibit Chemotaxis in Response to H. pylori-derived Peptide Hp(2-20) J. Immunol., June 15, 2004; 172(12): 7734 - 7743. [Abstract] [Full Text] [PDF]
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