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Departments of
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Cell Biology,
Pharmacology, and
Protein Expression, Human Genome Sciences, Inc., Rockville, MD 20850; and
§
Laboratory of Host Defenses, National Institute of Allergy and Infectious Diseases, 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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(MIP-1) on the surface of human monocytes
and dendritic cells, as inferred by its ability to compete for
[125I]MIP-1, but not for [125I]MIP-1ß
or [125I]monocyte chemotactic protein-1(MCP-1) binding to
intact cells. Based on calcium flux, MPIF-1 is an agonist on
CCR1-transfected HEK-293 cells, monocytes, and dendritic cells, but not
on CCR5-, CCR8-, or CX3CR1-transfected cells. The
inhibitory effect of guanosine
5'-O-(3-thio-triphosphate) (GTP-
S) or pertussis toxin
pretreatment on MPIF-1 binding and calcium mobilization, respectively,
indicates the involvement of G proteins in the interaction of MPIF-1
and its receptor(s). The increase in intracellular free calcium
concentration following MPIF-1 treatment is mainly due to the influx of
calcium from an extracellular pool. However, a portion of the
intracellular free calcium concentration is derived from a
phospholipase C inhibitor-sensitive intracellular pool. MPIF-1 induces
a rapid dose-dependent release of [3H]arachidonic acid
from monocytes that is dependent on extracellular calcium and is
blocked by phospholipase A2 (PLA2) inhibitors.
Furthermore, PLA2 activation is shown to be necessary for
filamentous actin formation in monocytes. Thus, the MPIF-1 signal
transduction pathway appears to include binding to CCR1; transduction
by G proteins; effector function by phospholipase C, protein kinase C,
calcium flux, and PLA2; and cytoskeletal
remodeling. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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All the known chemokines signal through two groups of seven
transmembrane receptors, CXCR and CCR, belonging to the rhodopsin-G
protein-coupled receptor superfamily (3). The extracellular binding
event results in the dissociation of the and ß subunits of the
G protein, initiating the activation of several different signal
transduction pathways, such as adenylate cyclase, protein kinase C
(PKC),2 tyrosine and
mitogen-activated protein kinases, and phospholipases A2,
C, and D (PLA2, PLC, and PLD, respectively) (3, 4).
Following activation, chemokine receptors rapidly become unresponsive
to further stimulation. This desensitization process is thought to be
caused by phosphorylation on residues present in the carboxyl tail of
the protein and receptor internalization (5, 6).
Monocytes and dendritic cells, which play an important role in the
pathogenesis of inflammation and in Ag presentation, are responsive to
chemokines. Monocytes express the chemokine receptors CCR1, CCR2, CCR5,
CCR8, CXCR2, and CXCR4 (7). Of the CC chemokines, MCP-1 is a potent
monocyte activator in vitro in that following binding to CCR2, it
induces intracellular calcium flux, actin polymerization, oxidative
burst, lysosomal enzyme, and cytokine release (8, 9). In comparison,
MIP-1, which binds CCR1 and CCR5, and MIP-1ß, which binds CCR5,
are weaker inducers of exocytosis and do not stimulate oxidative burst
and cytokine release (10, 11), suggesting perhaps functional
differences between the receptors (12).
The expression of chemokine receptors in dendritic cells depends on the developmental pathways and the stage of differentiation of the cells. Monocyte-derived dendritic cells express CCR1, CCR2, CCR5, CXCR1, CXCR2, and CXCR4 (7), while CD34+-derived dendritic cells also express CCR6 (13). CCR2 receptors in dendritic cells appear to have a substantially reduced functional activity, since calcium flux and chemotaxis induced by MCP-1 in this cell type are minimal (7). The effect of chemokines on dendritic cells seems to be restricted to chemotactic activity (7). Thus, chemokines might participate in the trafficking of dendritic cells, such as homing of progenitor cells to the periphery and migration of mature cells to lymphoid tissues, and in the recruitment of dendritic cells to inflamed mucosal sites (14, 15).
Myeloid progenitor inhibitor factor 1 (MPIF-1), a 99-amino acid CC
chemokine containing six cysteines, was recently identified in a cDNA
library derived from human aortic endothelium cells (16). The cysteines
at positions 33, 34, 57, and 73 are common to the other CC chemokines,
while those at positions 44 and 84 are not conserved. MPIF-1 is most
homologous to the MIP chemokine subgroup, with identities of 51 and
41% to MIP-1 and MIP-1ß, respectively, while having <40%
identity to the MCP subgroup. Although MPIF-1 has been characterized to
have chemotactic activity on monocytes and dendritic cells (16, 17) (B.
Nardelli, unpublished observations) and to inhibit colony formation of
bone marrow-derived low proliferative potential colony-forming cells
(16), little is known about its receptor specificity and mechanism of
action.
The aim of the present report is to investigate the receptor specificity and signal transduction pathway of a novel amino-terminal truncated form of MPIF-1 (amino acids 24–99), herein designated MPIF-1, in monocytes and dendritic cells. In addition, we compared the receptor specificity and the biological activities of MPIF-1 to the full-length MPIF-11–99 in monocytes. Using competitive binding of iodinated chemokines and receptor desensitization, we demonstrate that MPIF-1 binds and activates CCR1. Furthermore, we demonstrate that monocytes respond to MPIF-1 treatment by the activation of PLA2, a rapid increase in filamentous actin (F-actin) formation, and the release of [3H]arachidonic acid ([3H]AA).
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Granulocyte-macrophage CSF, IL-4, MIP-1, MIP-1ß, MCP-1, and
MCP-3 were purchased from PeproTech (Rocky Hill, NJ), and I-309,
RANTES, and full-length MPIF-1 were obtained from R&D Systems
(Minneapolis, MN). Iodinated MIP-1, MIP-1ß, and MCP-1 (2000
Ci/mmol) were purchased from Amersham (Arlington Heights, IL). MPIF-1
was iodinated with [125I]iodine, hydrogen peroxide, and
lactoperoxidase (2200 Ci/mmol; Amersham). Pertussis toxin, NDGA,
U73122, U73343, and staurosporine were purchased from Calbiochem (La
Jolla, CA). PMA and 4-BPB were purchased from Sigma (St. Louis,
MO). Manoalide was purchased from RBI (Natick, MA).
MPIF-1 expression and purification
The coding sequence of MPIF-1 was amplified from a human aortic endothelial library by PCR, during which unique restriction sites (SphI/HindIII) were introduced, thereby allowing the gene to be inserted into the expression vector pQE7. The resulted plasmid DNA was used to transform Escherichia coli M15 Rep4 host cells. The bacterial transformants were grown in Luria Bertoni medium containing ampicillin and kanamycin. Induction was performed at 1 mM isopropyl ß-[sca]d-thiogalactoside for 3 h. For large scale production, a semi-defined medium without antibiotics was used. MPIF-1 was produced as an insoluble protein deposited within inclusion bodies. MPIF-1 appears as a 9-kDa protein on reduced SDS-PAGE.
The recovery method for MPIF-1 used a series of procedures to separate the host contaminants through cell lysis, washes, and differential centrifugations to isolate inclusion bodies that contain partially purified MPIF-1. E. coli cell paste was suspended in a buffer containing 100 mM Tris (pH 7.4) and 25 mM EDTA. The cells were lysed by passing twice through a microfluidizer (Microfluidics, Newton, MA) at 6000–8000 psi. The lysed sample was mixed with NaCl to a final concentration of 0.5 M and then centrifuged at 7000 x g for 15 min. The resulting pellet was washed again with the same buffer plus 0.5 M NaCl and then centrifuged at 7000 x g for 15 min.
The partially purified inclusion bodies were then resuspended for 2–4 h at 20–25°C in 1.75 M guanidine hydrochloride containing 100 mM Tris (pH 7.4) and 25 mM EDTA. The sample was placed at 2–8°C overnight and subsequently centrifuged at 30,000 x g the next day. The supernatant (1.75 M guanidine hydrochloride extract) was mixed vigorously for 30 min at 4°C with 10 vol of a buffer containing 50 mM sodium acetate (pH 4.5), 125 mM sodium chloride, and 2 mM EDTA. Afterward the mixture was placed at 4°C without mixing for 1–48 h before the chromatographic purification steps described below.
The diluted MPIF-1 sample was clarified using a 0.22-µm pore size sterile filter (Pall Ultrafine Filtration, East Hills, NY). The MPIF-1 protein was then chromatographed over a strong cation exchange (POROS HS-50) column. The HS column was washed first with 6 column vol of a buffer containing 50 mM sodium acetate (pH 6.0) and 300 mM sodium chloride. The bound protein was eluted using 3–5 column vol of a stepwise gradient of 500, 750, 1000, and 1500 mM sodium chloride in 50 mM sodium acetate at pH 6.0.
The HS fraction eluted with 0.5 M sodium chloride was diluted 2-fold with water. The diluted sample was applied to a set of strong anion (POROS HQ-50) and weak cation (POROS CM-20) exchange columns in tandem mode. Both columns were washed with 50 mM sodium acetate (pH 6.0) and 150 mM sodium chloride. The carboxymethyl column was eluted with a 10- to 20-column vol linear gradient of 150-1000 mM NaCl. Fractions containing purified MPIF-1, as analyzed through SDS-PAGE and reverse phase HPLC, were combined. The carboxymethyl-purified MPIF-1 was loaded onto a size exclusion (Sephacryl S-100 HR) column for final polishing.
Cells
PBMC were purified from single donor Leukopacks (American Red Cross, Baltimore, MD) by centrifugation through a Histopaque gradient (Sigma). Monocytes were isolated from PBMC by counterflow centrifugal elutriation. Dendritic cells were generated by culturing elutriated monocytes in RPMI containing 10% FBS supplemented with granulocyte-macrophage CSF (50 ng/ml) and IL-4 (20 ng/ml) as previously described (18). Following 5–8 days in culture, the phenotype of the cells as identified by FACS analysis was >90% CD1a+, >90% MHC class II+, <10% CD14+, <5% CD3+, and <5% CD20+. HEK-293 cell lines expressing human CCR1 or human CCR5 and mouse pre-B cell lymphoma 4DE4 cells expressing human CCR8 have been previously described (19, 20, 21). 4DE4 pre-B cells were maintained in RPMI 1640 containing 10% FBS, 50 µM 2-ME, and 1 mg/ml G-418 at 37°C in 5% CO2 and 100% humidity. HEK-293 cell lines were maintained in DMEM with 10% FBS supplemented with 2 mg/ml G-418. THP-1 cells were obtained from American Type Culture Collection (Manassas, VA).
Whole cell binding assays
Competition binding studies were conducted using 1 x 106 monocytes, 5 x 105 dendritic cells, or 2 x 106 THP-1 cells/well. Cells were resuspended in binding buffer (Ham’s F-12 medium containing 0.5% BSA and 0.1% sodium azide) and transferred to a 96-well U-bottom plate (Costar, Cambridge, MA). The iodinated chemokine was then added in the absence or the presence of cold competitor chemokines in a total volume of 100 µl of binding buffer. The binding reaction was conducted for 90 min at 24°C with gentle agitation. Subsequently, the cells were pelleted by centrifugation at 1300 rpm for 5 min, and the supernatant was removed. The cells were washed with cold PBS (4°C) and then transferred to an opaque 96-well plate (Microfluor, Nunc, Newsbury Park, CA). Following a further centrifugation, the cells were resuspended in 100 µl of liquid scintillant and then counted.
Membrane binding assay
CCR1, CCR2b, and CCR5 membranes were purchased from Receptor
Biology (Beltsville, MD), and the assays were conducted as suggested by
the manufacturer. Briefly, CCR1-containing (25 µg), CCR2b-containing
(8.2 µg), or CCR5-containing (50 µg) membranes were incubated in a
96-well plate with [125I]MIP-1 (50 pM),
[125I]MCP-1 (55 pM), or [125I]MIP-1ß (500
pM), respectively, in the presence or the absence of cold competitor
chemokines in a total volume of 100 µl of binding buffer (50 mM HEPES
(pH 7.2), 5 mM MgCl2, 1 mM CaCl2, 0.5% BSA,
0.002% sodium azide, and protease inhibitors). Following an incubation
for 90 min at 24°C, the membranes were centrifuged at 2500 rpm for 10
min. The supernatant was decanted, 100 µl of binding buffer (4°C)
containing 0.5 M NaCl was added, and the membranes were transferred to
an opaque plate. Following two additional rinses, the membranes were
resuspended in 100 µl of scintillant and counted.
Measurement of changes in [Ca2+]i
Monocytes, dendritic cells, or THP-1 cells were resuspended in Ca2+ flux buffer (10 mM HEPES buffer (pH 7.4) containing 1 mM CaCl2, 2 mM MgSO4, and 5 mM glucose) and loaded with fura-2 (Molecular Probes, Eugene, OR; 0.2 nmol/106 cells) for 30 min at 37°C. The cells were then washed, resuspended in the labeling buffer, and treated with chemokine(s). Fura-2 fluorescence was measured on a fluorescence spectrophotometer (F-2000, Hitachi, San Jose, CA), and the ratio of the emission at 340 and 380 nm was determined. Calcium flux in cell lines transfected with CCR1, CCR5, and CCR8 was measured exactly as described previously (21).
Release of [3H]AA
The [3H]AA release assay was adapted from the protocol of Dumuis et al. (22). Freshly isolated monocytes were incubated overnight in RPMI containing 10% FBS and 1 µCi/ml of [3H]AA (205 Ci/mmol; Amersham). At the end of the incubation period, the cells were washed three times (10 min each) with RPMI containing 0.2% fatty acid-free BSA and then resuspended at the appropriate cell density in the same medium. Following the treatment period, the supernatant was removed, centrifuged, and counted. The remaining cell associated label was quantitated by solubilizing the cells in 0.2 M NaOH and counting the lysates.
Measurement of cAMP formation
The level of cAMP in the cultures of monocytes or dendritic cells was determined by ELISA according to the manufacturer’s instructions (Amersham).
Measurement of F-actin
Actin polymerization was analyzed as previously described (23). Monocytes (100-µl aliquots of 107 cells/ml) were preincubated with appropriate chemicals in HBSS containing 0.05% BSA and then treated with the chemokines at 37°C. The reaction was stopped by addition of fixing-staining buffer (137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, and 1.5 mM KH2PO4), pH 7.0, containing 8% (w/v) paraformaldehyde, 0.2 mg/ml lysophosphatidylcholine, and 0.3 µM Oregon Green 488 phalloidin (Molecular Probes). The cells were stained for 30 min at room temperature, washed, resuspended in PBS, and analyzed by FACScan (Becton Dickinson, San Jose, CA). The mean fluorescence intensity per cell was used as a measure of F-actin content per cell. Results are expressed as the relative fluorescence index, i.e., the ratio of the mean fluorescence intensity of chemokine-treated cells to the mean fluorescence intensity of control cells.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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bind to a shared site on monocytes and
dendritic cells
In competition binding studies using monocytes, MIP-1 and
MPIF-1 at a concentration of 40 or 1000 nM, respectively, displaced
50% of the bound [125I]MIP-1 (Fig. 1
A), while in dendritic or
THP-1 cells, 50% displacement was achieved with 1 nM MIP-1 or with
between 200–400 nM MPIF-1 (Fig. 1, D and G).
Concentrations of MIP-1 in the range of 10–100 nM produced maximal
displacement (70–90%) of [125I]MIP-1 in all three
cell types, while maximal displacement of [125I]MIP-1
was achieved with concentrations of MPIF-1 between 100-1000 nM.
Although specific [125I]MIP-1ß binding was observed in
all three cell types, MPIF-1, even at concentrations as high as 1000
nM, did not displace a significant amount of the bound
[125I]MIP-1ß (Fig. 1, B, E, and
H). The binding of [125I]MCP-1 on monocytes
was displaced by 50 or 85% with 0.7 or 100 nM MCP-1, respectively
(Fig. 1C). In contrast, in monocytes MPIF-1 did not displace
bound [125I]MCP-1 at any concentration tested. In
dendritic cells, 1000 nM MPIF-1 displaced approximately 20% of the
bound [125I]MCP-1, suggesting that the iodinated ligand
was binding to a low affinity site (Fig. 1, F and
I). In contrast, the displacement of
[125I]MCP-1 on THP-1 cells was nearly fully achieved with
0.5 nM MPIF-1. However, the maximal level of displacement achieved with
1000 nM MPIF-1 was 50%, in contrast to 90% competition in the
presence of 100 nM MCP-1. Based on the receptor specificity of
MIP-1, MIP-1ß, and MCP-1, the results of the above
cross-competition studies suggest that MPIF-1 binds to CCR1 on all
three cell types and possibly to CCR2 on THP-1 cells.
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To further delineate the receptor specificity of MPIF-1,
cross-competition studies were conducted on membranes from cell lines
expressing only one type of chemokine receptor. Using membranes from
HEK-293 cells expressing CCR1, the ability of MPIF-1 to compete with
[125I]MIP-1 binding was investigated (Fig. 2A). The competition curves
observed with increasing concentrations of MIP-1 or MPIF-1 were
similar, in that 50% displacement of the bound
[125I]MIP-1 was achieved with 50 nM MIP-1 or
MPIF-1. At a concentration of 500 nM, MIP-1 or MPIF-1 displaced
between 70–80% of the bound [125I]MIP-1. In
contrast, IL-8 in the same concentration range did not displace any
[125I]MIP-1 (data not shown). Since the N-terminal
region of ß-chemokines is known to be important in receptor binding
and activation, we compared the binding characteristics of MPIF-1 to
those of the full-length protein (1–99) on CCR1-containing membranes.
In competition binding experiments using [125I]MIP-1
and CCR1-containing membranes, the displacement curves generated with
the two proteins were not significantly different (Table I).
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B). The
binding selectivity of MPIF-1 for CCR5 was investigated by examining
the ability of MPIF-1 to displace either [125I]MIP-1
or [125I]MIP-1ß (Fig. 2C, right
or left panel, respectively). Although the unlabeled
chemokines effectively compete with their respective iodinated
counterparts for binding to CCR5, MPIF-1, even at a 2000-fold excess,
did not displace a significant amount of either bound
[125I]MIP-1 or [125I]MIP-1ß.
Furthermore, the full-length form of the protein did not displace a
significant amount of bound [125I]MIP-1 from CCR5
(Table I). These results demonstrate that the binding characteristics
of MPIF-124–99 and MPIF-11–99 are the same on
CCR1 and CCR5 and suggest that a shared MIP-1/MPIF-1 binding site on
monocytes and dendritic cells is CCR1.
When membrane binding experiments were conducted at 24 or 37°C, the
amount of [125I]MPIF-1 bound reached equilibrium within
30 min and remained stable for at least 90 min. Although the level of
binding at 4°C was initially lower than that observed at 24 or
37°C, equivalent levels of [125I]MPIF-1 binding were
observed at equilibrium (data not shown). Steady state binding
experiments were conducted to determine the affinity of
[125I]MPIF-1 for CCR1, and Fig. 3A depicts the binding
isotherm. Conversion of the data by Scatchard analysis revealed a
single affinity binding site with a Kd value of
745 pM (Fig. 3A, inset). The binding characteristics of
iodinated MPIF-1 were further investigated in cross-competition studies
using membranes from CCR1-transfected HEK-293 cells (Fig. 3B). In general, the competition curves for MPIF-1 and
MIP-1 were similar; however, MPIF-1 was somewhat more effective in
displacing the bound [125I]MPIF-1 than was MIP-1. At a
concentration of approximately 600 nM, MCP-3 displaced 50% of the
bound [125I]MPIF-1. MIP-1ß at the highest concentration
tested only displaced 20% of the bound [125I]MPIF-1 from
CCR1, consistent with its relatively weak affinity for CCR1.
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To test whether CCR1 is a functional receptor for MPIF-1, we
measured calcium flux in CCR1-transfected HEK-293 cells after
stimulation with MPIF-1. MPIF-1 induces a rapid calcium flux in these
cells, but not in untransfected HEK-293 cells or in cells expressing
CCR5 (Fig. 4A). Also, 4DE4
cells expressing the I-309 receptor (Fig. 4A) and HEK-293
cells expressing CX3CR1 (data not shown) failed to respond to MPIF-1.
The maximal receptor stimulation obtained with 100 nM MPIF-1 was
similar in magnitude to that obtained with the same concentration of
the other CCR1 agonists, RANTES, MIP-1, and MCP-3. Consistent with
this, MPIF-1 prestimulation was able to desensitize CCR1 to subsequent
stimulation with itself or RANTES, MIP-1, or MCP-3, whereas it had
no effect on the response of CCR5, CCR8, or CX3CR1 to known agonists.
The calcium response elicited by MPIF-1 treatment in CCR1-expressing
cells was dose dependent with a half-maximal effective concentration of
approximately 25 nM (Fig. 4B).
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To identify any potential difference in the biological
activities of MPIF-124–99 and the full-length protein, we
compared their abilities to induce cellular responses and their
patterns of receptor desensitization on monocytes. The two forms of the
protein had essentially identical potencies for inducing a
Ca2+ flux and they demonstrated equivalent activity in the
induction of F-actin polymerization (Table II; see below for details concerning
F-actin experiments). Furthermore, they induced equivalent patterns of
receptor desensitization (Table III).
When the cells were treated first with MIP-1 or MCP-3, the calcium
flux response to MPIF-1 was completely inhibited. In contrast, treating
the cells first with MCP-1 induced a partial desensitization of the
MPIF-1 response, while pretreatment with MIP1-ß, even at a
concentration as high as 100 nM, did not produce a significant
reduction in the response to MPIF-1. Furthermore, the pattern of
receptor desensitization induced by MPIF-1 was characterized in
dendritic cells and THP-1 (Table IV). In
general, the response pattern was equivalent to that observed in
monocytes. However, cross-desensitization between MPIF-1 and MCP-1 in
dendritic cells could not be studied, since MCP-1 does not induce
calcium flux in dendritic cells, as previously reported (7). The same
limitation was observed using MIP-1ß and THP-1 cells. The results
from the Ca2+ flux and receptor desensitization studies are
consistent with the data from the binding studies in demonstrating that
both forms of MPIF-1 bind to a subpopulation of receptors that also
bind MIP-1.
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The regulation of MPIF-1 binding to CCR1 by G proteins was tested
by preincubating membranes from CCR1-transfected HEK-293 cells with
GTP--S and then with [125I]MPIF-1. GTP--S inhibited
[125I]MPIF-1 binding in a concentration-dependent manner
with an IC50 of 10 nM (Fig. 5A). The involvement of G
proteins in the biological response to MPIF-1 was further identified by
testing the effect of pertussis toxin (PTX), which binds to and
inactivates Gi and Go proteins, on
the MPIF-1-associated change in [Ca2+]i.
Preincubation of monocytes with PTX completely inhibited the calcium
flux associated with MPIF-1 treatment (Fig. 5B).
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The increase in [Ca2+]i that is
associated with MPIF-1 receptor activation may be derived from
intracellular and/or extracellular calcium pools. To determine whether
the influx of extracellular calcium is the source of the
[Ca2+]i associated with the MPIF-1 response,
monocytes were loaded with fura-2, rinsed with flux buffer without
calcium and containing 2 mM EGTA, and subsequently stimulated with
MPIF-1 (Fig. 6B). Under these
conditions the [Ca2+]i response was less than
the response seen in the presence of extracellular calcium and in the
absence of EGTA (Fig. 6A). However, a residual peak of
[Ca2+]i was reproducibly observed when
monocytes were treated with MPIF-1, suggesting that a portion of the
calcium response is mobilized from an intracellular source(s). The
activation of PLC induces the formation of inositol triphosphate that
subsequently stimulates the release of calcium from intracellular
compartments. To ascertain whether MPIF-1 activates PLC, monocytes were
pretreated with U73122, a PLC inhibitor, and then with MPIF-1 in the
absence of extracellular calcium. Under these conditions, MPIF-1 did
not induce an increase in [Ca2+]i (Fig. 6C). In contrast, when the cells were pretreated with
U73343, a weakly acting analogue of U73122, and then with MPIF-1, there
was no reduction in the intracellular derived
[Ca2+]i peak (Fig. 6D).
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B). However, this
concentration of PMA had no effect on the basal levels of intracellular
calcium. The inhibitory effect of PMA was dose dependent, with an
IC50 value of approximately 5 nM (data not shown). To
ascertain whether the inhibitory effect of PMA requires the activation
of PKC, the cells were incubated first with staurosporine, a PKC
inhibitor, and then treated with PMA. Pretreatment with staurosporine
did not modulate the MPIF-1-induced [Ca2+]i
response in monocytes (Fig. 7C). However, staurosporine did
block the inhibitory effect of PMA on the MPIF-1-induced increase in
[Ca2+]i (Fig. 7D).
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The structural homology of CCR1 to neuropeptide Y and angiotensin
II receptors, both of which bind and activate adenylate cyclase, and
the regulatory effects that MIP-1 has on adenylate cyclase in MO7e
CFC cells suggest that ligands binding CCR1 may modulate cAMP in target
cells (24, 25). The possibility that MPIF-1 also affects adenylate
cyclase activity was investigated by treating monocytes, dendritic
cells, or THP-1 cells with MPIF-1 in a wide range of concentrations in
the presence or the absence of the phosphodiesterase inhibitor,
isobutylmethylxanthine, for time intervals ranging between 5–240 min.
No significant change in the level of cAMP was noted under any of the
conditions tested (data not shown).
To investigate whether MPIF-1 activates PLA2 in monocytes,
the cells were labeled overnight with [3H]AA and
subsequently treated with chemokines for different lengths of time.
Treatment with MPIF-1 (1000 ng/ml) produced a rapid time-dependent
increase in the amount of [3H]AA present in the medium
(Fig. 8A). The response was
dose dependent, with concentrations from 10–1000 ng/ml inducing a
linear increase in the released [3H]AA, and saturation
was achieved with 1000 ng/ml of MPIF-1 (Fig. 8B).
Pretreatment of the cells with manoalide, an inhibitor of
PLA2, blocked the release of [3H]AA following
MPIF-1 treatment. Furthermore, the stimulated release of
[3H]AA requires extracellular calcium, since the cells
were unresponsive to MPIF-1 in calcium-free medium containing 5 mM
EGTA.
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Chemokine-induced changes in cellular morphology, which ultimately
result in a chemotactic response, are produced in part by a
reorganization of the actin microfilament system. Quantitation of the
effects of MPIF-1 on F-actin formation are based on changes in the
amount of Oregon Green 488-labeled phalloidin bound to permeabilized
monocytes. A rapid dose-dependent increase in F-actin content was
observed when monocytes were treated with MPIF-1. The response reached
saturation within 10 s and then declined in a linear manner
approaching baseline levels after approximately 3 min (Fig. 9A). The polymerization of
F-actin induced by MPIF-1 was inhibited by 4-BPB and NDGA, inhibitors
of PLA2, and of 5-, 12-, and 15-lipoxygenase, respectively
(Fig. 9B). Furthermore, pretreating the cells with U73122
resulted in a near complete inhibition of the response.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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-S and PTX show that G protein(s) is involved in
MPIF-1 binding and receptor activation. In addition, we identified that
the source of the [Ca2+]i response associated
with MPIF-1 treatment is from both extracellular and PLC-inhibitor
sensitive intracellular pools. We also show that MPIF-1 induces a rapid
increase in the release of [3H]AA, demonstrating that
PLA2 activity is stimulated following treatment with this
CC chemokine. Activation of PLA2 is also required for the
rapid MPIF-1-dependent F-actin formation in monocytes.
The pattern of receptor binding with monocytes, dendritic cells, or
HEK-293 cells, expressing a single chemokine receptor, demonstrates
that MPIF-1 binds to a subpopulation of receptors used by MIP-1.
Cross-competition studies using membranes from cells expressing CCR1,
-2b, or -5, clearly show that MPIF-1 binds to CCR1 and that MPIF-1 and
MIP-1 have nearly equivalent affinities for this receptor. However,
the displacement curve for MPIF-1 and [125I]MIP-1
compared with that for MIP-1 was shifted to the right by as much as
2 logs in whole cell binding assays using monocytes, dendritic cells,
or THP-1 cells, suggesting that in these cell types MIP-1 has a
greater affinity for CCR1 compared with MPIF-1. Since the affinity
constants reported for [125I]MIP-1 and CCR1 are in the
range of those determined in this study for [125I]MPIF-1
and CCR1, an explanation for the differences in the displacement curves
based on a difference in affinities is not tenable (26). It is possible
that monocytes and dendritic cells may have additional extracellular
matrix components (e.g., glycosaminoglycans) that may affect MPIF-1
binding (27). Alternatively, since the receptor selectivity of MIP-1
is broader than that exhibited by MPIF-1, the displacement curve with
MPIF-1 would be less pronounced in cells expressing multiply chemokine
receptors.
The pattern of receptor selectivity of both forms of MPIF-1
characterized in binding assays is consistent with their receptor
specificity as identified on the basis of Ca2+ flux
studies. In experiments with HEK-293 cells transfected with CCR-1,
MPIF-1 induces a [Ca2+]i response, while
cells transfected with CCR5 are unresponsive. Furthermore, MPIF-1,
MIP-1, RANTES, and MCP-3 symmetrically cross-desensitize each other
on CCR1. The calcium desensitization reported in this study is
different from that previously reported by Forssmann et al. (17). In
the latter study, prestimulation of monocytes with MPIF-1 only
partially inhibits the response to a subsequent challenge with MIP-1
and completely desensitizes the cells toward MIP-1ß. On the basis of
our experiments involving calcium flux with HEK-293 cells expressing
CCR5, no interaction was detected between MPIF-1 and CCR5. Similarly,
neither full-length MPIF-11–99 nor
MPIF-124–99 demonstrated any affinity for CCR5 in binding
assays. In contrast, in displacement studies using CCR1-containing
membranes, MPIF-1 or the full-length protein produced an equivalent
displacement of [125I]MIP-1. Thus, the change in the
N-terminal sequence of MPIF-1 does not produce a change in its receptor
specificity or in its apparent affinity for CCR1.
The remodeling of membrane phospholipids by PLA2, PLC, and PLD is an essential step in the signal transduction pathways associated with the response of monocytes to CC chemokines and leads to the induction of diverse biological responses, including chemotaxis, activation of oxidative burst, and release of lysosomal enzymes (6). We have shown that both PLC and PLA2 are involved in the response of monocytes to MPIF-1. Approximately 30% of the [Ca2+]i response to MPIF-1 is derived from a PLC-sensitive intracellular pool. It is not known whether an increase in this intracellular pool of calcium has a preferential biological effect compared with the calcium derived from extracellular sources. However, pretreating the cells with the PLC inhibitor, U73122, blocks the F-actin response to MPIF-1, suggesting that the PLC-sensitive intracellular pool may preferentially regulate cell motility. In addition to the release of intracellular calcium, the activation of PLC results in the increase in PKC activity. Pretreatment of monocytes with PMA, an activator of PKC, results in the complete inhibition of the calcium response associated with MPIF-1 treatment. The effect of PMA is mediated through PKC, since the PKC inhibitor staurosporine blocks the response. However, staurosporine does not affect the calcium flux desensitization following MPIF-1 treatment. Activation of PKC has previously been shown to block the calcium response associated with MCP-1 treatment (28, 29). However, there are contradictory reports as to whether MCP-1 induces PLC activity in monocytes (28, 30). The mechanism of action of PKC is presently not known and may involve the phosphorylation of CCR1 and/or components of the MPIF-1 signal transduction pathway.
Treatment of monocytes with MPIF-1 induces a time- and dose-dependent
increase in [3H]AA release. This effect is due to the
activation of PLA2 and not to a nonspecific membrane
perturbation, since manoalide, a specific inhibitor of
PLA2, blocks the response, and the release of
[3H]AA is dependent on extracellular calcium. It has been
reported previously that treatment with MCP-1, MCP-3, RANTES, and
MIP-1, but not with MCP-2, induces the release of arachidonate from
monocytes (31). Studies involving treating cells with antisense
oligonucleotides to PLA2 have demonstrated that
PLA2 is necessary for the chemotaxis induced by chemokines,
but not by FMLP or C5a (32). A direct role for AA and its metabolites
has also been demonstrated to regulate the cell motility response
associated with other factors, e.g., epidermal growth factor or
fibroblast growth factor (33).
The formation of F-actin is an essential event in cellular adhesion, orientation, and motility. MPIF-1 induces a rapid concentration-dependent increase in the F-actin content in monocytes that is blocked by pretreating the cells with PLA2 inhibitors, e.g., 4-BPB. Furthermore, this PLA2 inhibitor blocks the MPIF-1-induced chemotaxis of dendritic cells (data not shown). Once formed, AA is further processed by both cyclo-oxygenases and lipoxygenases. The F-actin remodeling response observed following epidermal growth factor stimulation of A431 cells and neutrophils involves both 5-lipoxygenase and cyclo-oxygenase metabolites of AA (33, 34). The inhibition of 5-, 12-, and 15-lipoxygenases with NDGA results in the complete blockage of F-actin formation. The findings that some of the lipoxygenase metabolites of AA (e.g., 5-oxo-eicosatetraenoic acid and 5-hydroxyeicosatetraenoic acid) induce monocyte chemotaxis and AA release and act synergistically with CC chemokines indicate that they serve not only to amplify the response to CC chemokines in target cells but also as intracellular messengers affecting other cell types that potentially are unresponsive to the original chemokine (35).
The CCR1 expression pattern, based on immunofluorescence or RT-PCR, in
peripheral blood is restricted to monocytes, T lymphocytes, and NK
cells and in bone marrow and cord blood to CD34+ cells,
erythroblast, and erythroid progenitor cells (36, 37). On the basis of
the binding data and second messenger studies presented here for MPIF-1
demonstrating a selectivity for CCR1, other cells expressing CCR1 are
likely to be targets for this chemokine, and further comparison of the
biological activity of MPIF-1 to MIP-1 would be of interest.
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Acknowledgments |
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Footnotes |
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2 Abbreviations used in this paper: PKC, protein kinase C; PLA, phospholipase A; PLC, phospholipase C; MCP, monocyte chemotactic protein, MIP, macrophage inflammatory protein; MPIF-1, myeloid progenitor inhibitory factor-1; F-actin, filamentous actin; AA, arachidonic acid; [Ca2+]i, intracellular free calcium concentration; NDGA, nordihydroguaiaretic acid; 4-BPB, 4-bromophenacyl bromide; GTP-S, guanosine 5'-O-(3-thio-triphosphate); PTX, pertussis toxin.
Received for publication April 20, 1998. Accepted for publication September 2, 1998.
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