By
§
From the * Sir William Dunn School of Pathology, University of Oxford, Oxford OX1 3RE, United
Kingdom; Schering-Plough Laboratory for Immunological Research, Dardilly 69571, France; § Molecular Medicine Research Institute, Mountain View, California 94043; and
The DNAX
Research Institute, Palo Alto, California 94304
Dendritic cells initiate immune responses by ferrying antigen from the tissues to the lymphoid
organs for presentation to lymphocytes. Little is known about the molecular mechanisms underlying this migratory behavior. We have identified a chemokine receptor which appears to
be selectively expressed in human dendritic cells derived from CD34+ cord blood precursors,
but not in dendritic cells derived from peripheral blood monocytes. When stably expressed as a
recombinant protein in a variety of host cell backgrounds, the receptor shows a strong interaction with only one chemokine among 25 tested: the recently reported CC chemokine macrophage inflammatory protein 3. Thus, we have designated this receptor as the CC chemokine receptor 6. The cloning and characterization of a dendritic cell CC chemokine receptor
suggests a role for chemokines in the control of the migration of dendritic cells and the regulation of dendritic cell function in immunity and infection.
Dendritic cells (DCs)1 are central to antigen-specific immunity in vertebrates. They act in vivo as immune
potentiating cells, sampling the antigenic environment and
presenting those antigens to effector lymphocytes (for reviews
see references 1 and 2). DCs are found at many sites in the
body, particularly at or near surface areas where antigen
contact is likely, and in the lymphoid organs where they
form close associations with T cells. Migration is an integral
part of DC function; precursor cells must migrate from the
bone marrow to resident sites in the tissue. After exposure to antigen or inflammatory stimuli, many DCs migrate from
their resident sites to the secondary lymphoid organs (1).
Much of how DCs achieve their migratory functions remains
a mystery. Recent attention has focused on the chemokine
system, a multipartite superfamily of chemoattractant cytokines that induce the directed migration of leukocytes and
other cells, and on the superfamily of G protein-coupled,
seven transmembrane receptor proteins through which chemokines act (for reviews see references 5). Chemokines are
involved not only in immune cell trafficking and inflammation, but are also central to infectious disease processes. For
example, certain chemokines have been shown to be endogenous inhibitors of HIV-1 entry (9), and some chemokine
receptors are `cofactors' or `second receptors' for HIV binding
and fusion with target cells (10).
The understanding of how chemokines might regulate
the migration of DCs is still rather sparse, though a few reports clearly demonstrate that certain CC chemokines are
indeed chemoattractants for some types of DC in vitro (16,
17). It has been also demonstrated that HIV-1 can infect
DCs in vitro through interactions with CC chemokine receptor (CCR)5, CXC chemokine receptor (CXCR)4, and probably another chemokine receptor (18). To address some
of the questions surrounding the control of DC function at
the molecular level, we sought to identify new elements of the
chemokine system in these cells. Here, we report on the isolation of a new chemokine receptor, CCR6, which is highly
expressed in CD34+ cell-derived DCs, but not in monocyte-derived DCs, and its interaction with a recently described CC chemokine, macrophage inflammatory protein
(MIP)-3 PCR Screening of cDNA Libraries.
cDNA plasmid libraries from
DCs, eosinophils, and T cells were used as templates in PCR reactions as follows: 50 µl total volume in 0.2-ml tubes with 5 µM
degenerate primers second intracellular loop (IC2) sense (5 Analysis of Gene Expression Distribution.
Northern blot analysis was performed using a multiple tissue Northern blot purchased
from Clontech (Palo Alto, CA). Where limited amounts of messenger RNA (mRNA) precluded direct Northern analysis, expression was assessed on Southern blots of cDNA libraries. Libraries were constructed from polyA+ RNA selected with
oligotex beads (Qiagen, Chatsworth, CA). 2 µg of mRNA was
primed by oligo dT (Superscript cDNA synthesis kit; GIBCO
BRL), and the quality of the resulting cDNA was evaluated by
three criteria: (a) a size range of cDNA from the first strand synthesis ranging from >0.5 to 5 kb on alkaline gel analysis, (b) independent clones numbering greater than 106/100 ng of unamplified cDNA, and (c) a high proportion of full-length clones with
low levels of genomic or ribosomal RNA contamination (<5%),
as assessed by random sequence analysis of each library. For
Southern analysis, large scale plasmid DNA preparations of amplified libraries were done using a Giga prep (Qiagen). 5 µg from
the primary amplified cDNA library was digested with NotI and
SalI (Boehringer Mannheim, Indianapolis, IN) to release the inserts, run on a 1% agarose gel, and transferred to a nylon membrane (Schleicher & Schuell, Keene, NH). Blots were probed
with a 32P-labeled BN-1 cDNA fragment under high stringency
hybridization and wash conditions. The resultant hybridization
patterns reflect the different sized cDNA fragments in the libraries
and their relative abundance. To confirm that the restricted distribution was reflective of unamplified mRNA, semiquantitiative
PCR was performed on total RNA isolated directly from monocytes, monocyte-derived DCs, and CD34+ cell-derived DCs. The
amplification pattern mirrored the hybridization patterns seen in
the library Southern blots.
Chromosomal Localization.
Genomic DNA (400 ng) from a
panel of 20 human-rodent somatic cell hybrids cell lines (Biosmap; BIOS Labs., New Haven, CT) was used as a template in a
50-µl PCR reaction. The primer pairs were: BN1, 5 Receptor-Ligand Binding and Signaling Analyses.
Full-length BN-1
expression constructs were first made in pcDNA3 (Invitrogen,
San Diego, CA) and used to generated stable transfectants in human embryonic kidney 293 (HEK293) cells as described (20). To
control for levels of cell-surface expression of the receptor, subsequent constructs were made in which the M2 "Flag" epitope was
attached to the NH2-terminal portion of the receptor. Anti-M2
antibody was then used to detect cell-surface BN-1-encoded
polypeptide and to sort the highest expressing cells for stable
propagation. Intracellular calcium signaling analyses were performed as described (21). Equlibrium binding was done using a
standard filtration protocol using 106 cells incubated with 0.1 nM
[125I]MIP-3 Degenerate reverse transcription PCR was used to identify
novel chemokine receptor-like sequences from several cell
types: monocytes, cultured T cells, eosinophils, and DCs. A
combination of primer pairs was used representing homologous regions within the family of known chemokine receptors and other seven transmembrane-spanning receptors
associated with cell motility. Primer pairs from regions encoding TM2 and TM7, as well as the IC2, were used on
substrate mRNA or cDNA from the cells listed above. Of
these, primer pair IC2 sense and TM7 antisense generated
from DCs, and to a lesser extent eosinophils, a 550-bp
PCR product whose sequence was at that time not present
in any of the available databases. The 550-bp cDNA representing the IC2-TM7 segment were then used to isolate from a DC library a larger cDNA clone, designated `Barney' or BN-1, encoding the entire presumptive open reading
frame (ORF) of a novel chemokine receptor-like protein.
The predicted protein sequence encoded by the BN-1
ORF is shown in Fig. 1, aligned with human CXCR2
(formerly known as IL-8RB) and human CCR4. During
further characterization of the function of the BN-1 protein, the same sequence was subsequently deposited as an
"orphan" TM7 receptor in the EMBL/GenBank/DDBJ
database as accession No. U68030, and was reported as a
potential chemokine-like orphan receptor sequence of unknown function (22, 23). Multiple sequence alignment of
the protein encoded by BN-1 with other human chemokine receptor sequences showed amino acid identity with
CXCR1 and CXCR2 of ~38%, but this is only slightly
more identity than to the closest human CC chemokine
receptor (CCR4, ~36%). The sequence alignment of Fig.
1 shows that parts of the BN-1-encoded ORF share extensive homology with CXC receptors, but not CC receptors
(e.g., within TM3), whereas other parts of the BN-1 ORF
show closer homology to CCR4 (e.g., the intracellular
TM3-TM4 loop). Phylogenetic depictions of the evolutionary relatedness of the various chemokine receptors show
BN-1 to be on a branch containing CXCR1, CXCR2,
and CXCR4, and distinct from the branch containing the
previously described human CC chemokine receptors
CCR1-CCR5 (not shown).
We analyzed a
panel of human-rodent somatic cell hybrids to ascertain the
location of the human BN-1 gene. PCR using two pairs of
BN-1-specific oligonucleotides consistently yielded a PCR
product in only those hybrids containing human chromosome 6 (Fig. 2 A). Positive controls using PCR primers
against a known human chromosome 6 gene, human TNF,
confirmed the hybrid cell line karyotyping (Fig. 2 B), and
localization of BN-1 on human chromosome 6 was confirmed by PCR typing of chromosome 6 deletion and
translocation hybrids (not shown). We next used a radiation hybrid panel (Fig. 2 legend) to unambiguously localize
the BN-1 gene to 6q26-27, consistent with the fluorescent
in situ hybridization analysis of Liao, et al. (23). This constitutes a new locus for chemokine receptors; the CC
chemokine receptors CCR1-CCR5 are closely linked on
chromosome 3p21, the CXC receptors CXCR1 and
CXCR2 map to chromosome 2q35, and CXCR4 maps to
chromosome 2q21.
Various direct and indirect
approaches were used to assess the distribution of BN-1
mRNA. A Northern blot containing mRNA from a variety of organs and tissues showed an ~3.5-kb message predominantly in spleen, and to a lesser extent in thymus, testis, small intestine, and peripheral blood (Fig. 3 A, top left). In addition, the spleen showed two smaller transcripts of ~2.7
and 1.7 kb. mRNAs from various lymphoid and hematopoietic cell lines (TF-1, Jurkat, MRC5, JY, and U937; Fig 3
A, top right) were negative for BN-1 expression. For cell
lines and tissues where limited amounts of mRNA precluded direct Northern analysis, expression was determined
by the extent of hybridization among the gel-fractionated population of cDNA inserts from libraries made from those
cells. BN-1 expression was examined first in 19 cDNA libraries made from various cells of lymphoid lineage (Fig. 3
A, bottom). BN-1 cDNA was present in one library made
from resting PBMCs, consistent with the observation of
Zaballos et al. for expression of the CKR-L3 orphan cDNA
in CD4 and CD8 cells (22), and of the STRL22 orphan in
PBLs (23). Interestingly, however, a matched PBMC library
made after the cells were activated with anti-CD3 antibody and PMA showed no BN-1 cDNAs, which was further
notable by their absence from virtually every other library
made from T cell lines and clones (in various states of activation and anergy), pooled B cells, and NK cells (Fig. 3 A,
bottom). Resting human splenocytes contain BN-1 cDNA
(Fig. 3 A, bottom, Splen), but a matched library made after
activation of splenocytes with anti-CD40 and IL-4 (Splen Act) showed diminished levels of BN-1 cDNA. Thus it appears that BN-1 may not be abundantly expressed in the
lymphoid lineage, or that its expression is downregulated
with cellular activation or growth in lymphocyte cultures.
Since the original BN-1 PCR product was generated
from a library derived from human DCs, we assessed the
distribution of the receptor among these cells. There are
two principle methods by which DCs can be generated in
vitro: (a) stimulation of peripheral monocyte progenitors
by prolonged exposure of these cells to GM-CSF and IL-4
(24, 25), and (b) by culture of CD34+ precursor cells from
bone marrow or cord blood with GM-CSF and TNF. Resultant DCs can be tracked via expression of differentiation markers such as CD1a and CD86, and purified by sorting
for one or more of these markers. DCs generated by both
procedures take on the typical veiled morphology and
achieve antigen presentation capability (26, 27).
A striking distribution of BN-1 cDNA was seen in a
panel of libraries made from monocytes and two types of in
vitro-derived dendritic cells, suggesting marked differences
between monocyte- and CD34-derived DCs. BN-1 was
not present in any of the libraries derived from purified
elutriated monocytes, and there were few if any BN-1
cDNAs in the libraries prepared from monocyte-derived DCs (Fig. 3 B, top). By contrast, there is clear expression of BN-1 as the CD34+ cells develop into differentiated DCs
(Fig. 3 B, top). Although little signal is seen when these cultures are 30% CD1a+ (at about day 6), abundant message is
present by the time the cells are 70% CD1a+ (about day
12). Interestingly, when 70% CD1a+ cultures are stimulated for 1-6 h with PMA and ionomycin (70% CD1a+ act
6 h), much less BN-1 cDNA is found in the libraries subsequently made from those cells (Fig. 3 B, top). BN-1 is most
abundantly represented in libraries made from DC cultures
derived from cells which were sorted on the basis of CD1a
expression (95% CD1a; Langerhans-like), and less well in
those DCs that were derived from CD14+-sorted cells
(dermal/interstitial-like) (Fig. 3 B, top; reference 27). To
control for the quality of monocyte and monocyte-derived DC libraries, we probed with other markers, including the
CC chemokine human CCF-18/MIP-1 The chemokine literature is replete with cDNA sequences
encoding orphan chemokine receptors of unknown ligand
specificity and unknown function. To assess the ligand-binding specificity of the putative chemokine receptor encoded
by BN-1, we constructed expression plasmids (pFlagBN-1)
encoding a receptor with an added NH2-terminal Flag
epitope. This allowed for detection and selection of the
most highly expressing stable transfectants using an anti-Flag monoclonal antibody. To decrease the chance of a
species- or lineage-specific G protein-dependent response,
we also tested BN-1 expression constructs in the following
different types of cells from different species: human embryonic kidney 293 (HEK293), murine 3T3 fibroblasts, and chinese hamster ovary (CHO) cells. Stable transfectants were
generated from each of these cell lines; for some, cells were
sorted by anti-Flag antibody for high expression and further
propagation, an example of which is shown in Fig. 4 A.
A panel of purified chemokine ligands was then used to
challenge the various stable transfectants expressing the
BN-1 gene product. Calcium mobilization in the cytoplasm of BN-1 transfectants was used to assess receptor engagement and functional coupling to G proteins, as has
been established for other chemokine receptors (21). We
tested a panel of 25 purified recombinant or synthetic chemokines (listed in Fig. 4 B) including all of the standard CXC, C, and traditional CC chemokines, as well as the
CX3C chemokine Fractalkine (20). With one exception, all
chemokines were negative for calcium mobilization in recombinant BN-1-bearing cells of all backgrounds. One
chemokine, however, the recently described CC chemokine MIP-3 We report the identification of a cDNA encoding a
chemokine receptor abundantly expressed in certain types
of cultured human DC populations, and the characterization of the protein that it encodes. This receptor, designated here as CCR6, is notable in that of the over two
dozen purified chemokines tested, it appears to interact
only with the newly reported CC chemokine MIP-3 CCR6 is expressed most highly in DCs derived from
CD34+ cells, but not in monocyte-derived DCs. We have
also noted CCR6 expression in resting PBMCs, consistent
with reports of the presence of transcripts for the orphan
clones CKR-L3 and STRL22 (22, 23). Those reports, however, did not examine DC populations, nor did they
identify any ligands which bound the orphan cDNA gene
product. The robust but differential patterns of CCR6 suggest that it plays a role in the function of certain types of
DCs. At least three populations of cells with dendritic morphology have been reported in peripheral blood (29, 30).
Each may represent discrete maturational and functional
stages; the possibility has been discussed that the CD34-derived DCs in vitro are enriched for Langerhans-like cells. It will be therefore be of interest to assess whether CCR6
will prove to be DC lineage marker in vivo. It is known
that of the different DC populations identified in the periphery, only one appears to efficiently support HIV-1 infection (30). DCs have been proposed to act in a variety of
sites as a reservoir for HIV-1 (31), although most efficient
infection seems to occur in cocultures of DC and T cells.
The virus can clearly infect DCs alone, and can use the
chemokine receptors CXCR4 and CCR5 as infection cofactors (18). Additionally, some infection of DC from
CCR5-deficient individuals by M-tropic HIV has been
observed (18), suggesting an additional coreceptor on these
cells. Although we do not yet know how CCR6 expression relates to the different types of DCs in vivo, it may be
a candidate for an infection cofactor for strains of HIV that
infect DC (32).
It has been shown that monocyte-derived DCs and
CD34+ umbilical cord blood-derived DCs are chemotactically responsive to the CC chemokines RANTES, MIP-1 In summary, the identification of CCR6 as a provisionally specific receptor for the CC chemokine MIP-3 (19).
GATCGNTAGCTNGCNATNGTNCA T/C GC) and transmembrane (TM)7 antisense (5
GCATANATNANNGG G/A T
C/T NAN G/A CAGCAGTG) in buffer containing 20 mM
NH4S04, 75 mM Tris HCl, pH 9.0, at 25°C, 0.01% Tween 20, 2 mM MgCl2, 200 mM deoxynucleotide triphosphates and 2.5 units Taq DNA polymerase (ABT, London, UK). "Touchdown" PCR was performed as follows: 15 cycles of 94°C for 1 min, 65- 50°C annealing for 1 min decreasing by 1° per cycle, and at 72°C for 2 min followed by 20 cycles of 94°C for 30 s, 50°C for 1 min, and 72°C for 2 min. PCR products (550 bp) were purified after agarose gel electrophoresis by absorption to glass beads (Qiaex II;
Qiagen GmbH, Hilden, Germany) and cloned via Taq DNA
polymerase A overhangs into the T-tailed vector pTAg1 (R&D
Sys. Inc., Abingdon, UK). Recombinant plasmids were identified
by colony PCR and restriction mapping, and double-strand sequencing was performed using flanking M13 and T7 primers.
Full-length BN-1 cDNA sequence was obtained by PCR amplification using BN-1-specific oligonucleotides and M13 forward
and reverse primers to PCR amplify BN-1 sequences from human DC cDNA libraries constructed in the cloning vector pSPORT (GIBCO BRL, Gaithersburg, MD). In addition, radiolabeled probes generated from the PCR products were used to
isolate full-length BN-1 clones by colony hybridization.
GACCGGTACATCGCCATTGTACAGGC; BN2, 5
CTGAACTTC-
TGCCCAATAAAAGCGTAG; BN3, 5
GTACAAGTCCTCAGGCTTCTCCTG; and BN4, 5
TGCATAACATCTATGAGTATGTTTCAC. Human TNF-
promoter primers TNF1:
5
CAAACACAGGCCTCAGGACTC and TNF2: 5
AGGGAGCGTCTGCTGGCTG were a gift of Dominic Kwiatkowski. Genomic DNA from a panel of 16 somatic cell hybrids containing deletions and translocations of human chromosome 6 were a
gift of Dr. J.M. Boyle (Paterson Institute, Manchester, UK). Genomic DNAs of the radiation hybrid mapping panel Genebridge
4 were obtained from the Medical Research Council HGMP
Resource Centre (Cambridge, UK).
(custom labeled by Amersham, Arlington Heights,
IL) in the presence increasing amounts of unlabeled MIP-3
competitor. Reactions were incubated for 2 h at 22°C before being aspirated onto GF/C filters, washed, and measured by scintillation counting. Data were analyzed using IgorPro software
(WaveMetrics, Lake Oswego, Oswego, OR).
Cloning of Novel Chemokine Receptor Candidate cDNAs.
Fig. 1.
Sequence homology of the BN-1-coding region with other
chemokine receptors. The deduced 374-amino acid sequence encoded by the BN-1 cDNA was compared to other human and viral chemokine receptors using the multiple sequence alignment program Pileup of the Wisconsin EGCG DNA analysis software package. Shown here is the BN-1
amino acid sequence aligned with the human CXCR2 and CCR4. The
positions of the hydrophobic membrane spanning regions TM1-TM7 are
indicated by bars above the sequence. Amino acids identical between
BN-1 and either CXCR2 or CCR4 are boxed.
[View Larger Version of this Image (62K GIF file)]
Fig. 2.
The BN-1 gene maps to chromosome 6. (A) PCR analysis of
hamster, mouse, human and rodent-human hybrid cell line genomic
DNAs (samples 1-20 Biosmap; BIOS Labs., New Haven, CT) using BN-1-specific primers BN1 and BN2. The lane marked Blank is the result of
PCR in the absence of template, lanes M1 and M2 contain DNA molecular weight markers. The same genomic DNA samples gave positive
PCR signals of the expected size with a second pair of BN-1-specific PCR primers, BN3 and BN4 (see Materials and Methods). (B) The result
of PCR analysis of the same genomic DNA samples with human TNF-
promoter primers. The assignment of the BN-1 gene to chromosome 6 was confirmed by PCR analysis of a panel of chromosome 6 deletion and
translocation hybrids (data not shown). PCR analysis of the Genebridge 4 radiation hybrid panel with BN1 and BN2 primers gave the following data
vector: 12202021002210000101200020000000000000000112010000100 0010200011000020210100001020010100000001. The BN-1 data vector was compared to the WICGR human genome radiation hybrid map
using the Whitehead Institute/Massachusetts Institute of Technology Center for Genome Research automapper version 1.0 (http//:www-genome.wi.mit.edu/). The BN-1 STS was placed on chromosome 6, 3.36 centiRay from the framework STS D6S1008 with a lod score >3.0.
[View Larger Version of this Image (94K GIF file)]
Fig. 3.
Analysis of BN-1 gene expression. (A) Northern blot analysis
of BN-1 expression in RNA prepared from the various human tissues and
cell lines (top); sizes of RNA markers (in kilobases) are indicated in the left
margin. (Bottom) A Southern blot analysis of BN-1 in cDNA libraries prepared from the human PBL and primary cell T, B, and NK cell cultures.
PBMC, human peripheral blood mononuclear cells; PBMC Act, the same
PBMC stimulated with anti-CD3 and PMA for 2, 6, and 12 h and
pooled; Mot 72 and Mot 81, human Th0 T cell clones. Act, Stimulation
with anti-CD3 and anti-CD28 for 2, 6, and 12 h and pooled; and Anergic,
stimulation with a specific peptide rendering the cells nonresponsive to
antigen stimulation. HY06 and HY93 are human Th1 and Th2 T cell
clones, respectively, with activation and anergy treatments as above except for the peptide specificity; B cell pool, a collection of EBV cell lines;
Splen, total human splenocytes, resting; Splen Act, the same population
stimulated with anti-CD40 and IL-4 for 2, 6, and 16 h and pooled; NK
pool, a pool of primary NK cell clones; NK Act 6h, the same pool stimulated 6 h with PMA and ionomycin; NK B1, a single primary human NK
cell clone; NK Act 6h, that clone activated as above. Probing replicate
blots with human actin cDNA gave readily detectable ~2.0-kb species
in all lanes (data not shown). References upon request. (B) Southern blot
of BN-1 distribution (top) in cDNA libraries made from monocytes and
DCs. U937, human monocyte cell line. Human elutriated monocytes
have been stimulated as follows: LPS/IFN
, cultured in the presence of
these activators and blocking antibodies for IL-10 for 1, 2, 6, 12, and 24 h
and pooled; LPS/IFN
/IL10, the same pool without blocking antibodies
to IL-10; LPS 1 h and LPS 6 h, monocytes stimulated for 1 and 6 h with
LPS, respectively; CD34-derived DC, 30 and 70% DCs are CD1a+ DCs
derived from CD34+ human cord blood stem cells by growth in GM-CSF
and TNF-
, with the percent CD1a+ cells determined by FACs® over
time in culture. The cultures were 70% CD1a+ after 12 d. 70% CD1a+
act 1 h and act 6 h, the same cultured DC stimulated with PMA and ionomycin for 1 and 6 h, respectively. Mono-derived DC, DCs derived from
human elutriated monocytes by growth in GM-CSF and IL-4 for 5 and
10 d as listed; GM/IL4/LPS act, 10-d cultures stimulated for 6 h with
LPS; GM/IL4/IL1+TNF, are 10-d monocyte-derived DCs stimulated
with IL-1
and TNF-
for 4 and 16 h and pooled. CD34-derived DC,
sorted, DCs derived from CD34 cells as described that were then sorted on
the basis on the cell surface expression of the markers listed: 95% CD1a+,
DC derived from CD1a-sorted cells (Langerhans-like); CD1a+/CD14+, DC derived from CD14-sorted cells (dermal/interstitial). (Bottom) The same
blot stripped and reprobed with the human CCF-18/MIP-1
-chemokine. In all cases, the ladder effect represents different sizes of cDNA inserts in the
library ranging from ~0.6 to 3.5 kbp for BN-1. The two predominant CCF-18/MIP-1
cDNAS are ~0.7 and 1.3 kbp. (C) PCR analysis of unamplified mRNA from DC to confirm BN-1 distribution. Lane 1, CD34+ cord blood cells cultured 12 d in GM-CSF and TNF-
; lane 2, CD34-derived DCs stimulated with PMA and ionomycin; lane 3, CD34-derived DC purified by FACS® sorting for 98% CD1a+ expression; lanes 4-8, various cultures
of monocyte-derived DCs (cultured in GM-CSF + IL-4 for 8 d) under conditions of stimulation as shown; + Ctl, positive control amplification using 1 pg BN-1 plasmid as starting substrate;
Ctl, same reaction with identical reagents in the absence of the plasmid substrate. Elutriated monocytes (not
shown) were also negative. Each lane is representative of at least three independent experiments.
[View Larger Versions of these Images (81 + 88 + 42K GIF file)]
(28). Strikingly,
the cDNA distribution pattern for this chemokine is almost
directly inverse of that for BN-1 (Fig. 3 B, bottom). Finally,
PCR performed directly on primary, unamplified poly A+
mRNA harvested from monocyte- and CD34-derived DCs
yielded amplification products whose pattern mirrored the
hybridizations seen in the library southern blots (Fig. 3 C).
Taken together, the distribution data suggest that BN-1
seems to be expressed most highly in a mature phenotype
of DCs derived from the differentiation of CD34+ cells,
and is not well expressed in DCs generated from monocytes.
Fig. 4.
Chemokine specificity of the receptor encoded by
BN-1. (A) A sorted population
of transfected CHO cells stably
expressing BN-1 protein containing an NH2-terminal Flag
epitope (CHO-FlagBN-1) showing intensity of anti-Flag mAb
staining relative to wild-type CHO cells. (B) Panel of purified
recombinant or synthetic chemokines assayed for intracellular
calcium mobilizing activity in
various BN-1 transfectants. At
100 nM final concentration indicates no detectable intracellular calcium mobilization in any
of the transfectants (n >3),
whereas ++ denotes a robust
response in all transfectants
(CHO, 3T3, HEK293) bearing
BN-1. (C) Dose response of
MIP-3
in the induction of intracellular calcium in CHO-FlagBN-1 cells. CHO wild-type
cells are shown as a control. (D)
Binding and homologous competition of radiolabeled MIP-3
to BN-1-bearing cells. (Inset)
Scatchard transformation of the
binding data to reveal a Kd of
~0.1 nM.
[View Larger Version of this Image (21K GIF file)]
, showed a robust calcium signal, whereas the wild-type and mock-transfected cells were unresponsive.
MIP-3
exhibited a dose-dependent response, reaching a
plateau at ~10 nM (Fig. 4 C). MIP-3
was positive in every test on the stable transfectants in all three cellular backgrounds and also in transfectants expressing the native form
of the receptor (without the NH2-terminal flag), as well as
in a variety of transiently transfected cells of different types
(not shown). We did not detect MIP-3
activity on other
chemokine receptors tested (CCR1-CCR5, and CXCR1
and 2 [not shown]). Competition binding and dissociation
experiments were also performed. Homologous competition of radioloableled MIP-3
with unlabeled ligand showed
clear dissociation; Scatchard transformation revealed a dissociation constant (Kd) of ~0.1 nM (Fig. 4 D). From the
signaling and binding data, we therefore concluded that the
BN-1 gene product is a receptor for the CC chemokine
MIP-3
, hence its designation as CCR6.
.
, and monocyte chemotactic protein 3 (16, 17). The
chemotactic response of DCs to MIP-3
, however, has not
been reported. Indeed, much remains to be determined as to the properties of MIP-3
. Recently identified as an orphan chemokine of unknown function (19), it has also
been described as LARC (liver activation-related chemokine) by Hieshima et al. (33). Expressed sequence tags containing parts of the MIP-3
-coding region were generated
from sequence analysis of fetal lung, liver, and pancreatic islet cDNA libraries (19, 33). Although a CC chemokine,
MIP-3
is widely diverged from other CC chemokines, showing only 20-28% identity with other members of the
CC family and being encoded on chromosome 2 rather
than chromosome 17 (33). The chemotactic properties of
MIP-3
, especially as mediated through CCR6, are currently being studied. LARC protein was shown to induce chemotaxis of T cells, albeit less potently and efficiently
than RANTES (33). This would be consistent with CCR6
expression in T cells as noted here and for CKR-L3 (22),
but the existence of other MIP-3
receptors on lymphocytes cannot be excluded. Our experiments suggest that
levels of CCR6 expression in other cells are low relative to
the levels seen in DCs, and it is notable that CCR6 so far
exhibits specificity only for MIP-3
among the extensive
panel of chemokines tested. The specificity of CCR6 binding is supported by the evidence of Baba et al., whose study appeared while the present manuscript was under revision,
reporting a receptor of identical sequence, GPR-CY4, that
binds specifically to LARC/MIP-3
(34). Again, that study
did not examine dendritic cell distribution or function.
defines
another piece in the puzzle of chemokine-chemokine receptor interactions. Detailed analysis of CCR6 and its CC
chemokine ligand may shed new light on the control of DC
migration and function, providing a potential therapeutic
avenue for treatment of a wide range of pathologies including
HIV infection, autoimmunity, and transplant rejection.
Address correspondence to Thomas J. Schall, Molecular Medicine Research Institute, 325 E. Middlefield Rd., Mountain View, CA 94043. Phone: 415-237-7442; FAX: 415-237-7455; E-mail: tschall{at}mmrx.org
Received for publication 24 April 1997 and in revised form 24 June 1997.
DNAX is supported by Schering-Plough; work in the laboratory of S. Gordon is supported by the United Kingdom Medical Research Council; D.R. Greaves is supported by Glaxo Wellcome.We thank Dr. K. Bacon for help and advice, and Drs. J.M. Boyle and M.J. Greaves for gifts of human chromosome 6 hybrid DNAs and advice on human gene mapping studies. We are grateful to Jasmine and Shona for stimulating discussions regarding cell motility in the early stages of this project.
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