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Originally published online as doi:10.1189/jlb.0303116 on August 1, 2003

Published online before print August 1, 2003
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(Journal of Leukocyte Biology. 2003;74:868-879.)
© 2003 by Society for Leukocyte Biology

Prostaglandin E2 inhibits production of the inflammatory chemokines CCL3 and CCL4 in dendritic cells

Huie Jing, Evros Vassiliou and Doina Ganea1

Department of Biological Sciences, Rutgers University, Newark, NJ 07102

1Correspondence: Rutgers University, Dept. of Biological Sciences, 101 Warren St., Newark, NJ 07102. E-mail: dganea{at}andromeda.rutgers.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Dendritic cells bridge innate and adaptive immunity and participate in both responses. Upon capture of pathogens, dendritic cells release inflammatory cytokines and chemokines, attracting other immune cells to the infection site. Anti-inflammatory cytokines, glucocorticoids, anti-inflammatory neuropeptides, and lipid mediators such as prostaglandin E2 (PGE2) limit and control the inflammatory response. In this study we report that exogenous PGE2 inhibits CCL3 (MIP-1{alpha}) and CCL4 (MIP-1ß) expression and release from dendritic cells stimulated with either lipopolysaccharide (LPS), a TLR4 ligand, or peptidoglycan, a TLR2 ligand. The inhibition is dose-dependent and occurs at both the mRNA and protein levels. The inhibitory effect is mediated through EP2 and EP4 receptors and requires the presence of PGE2 at the time of LPS stimulation. Intraperitoneal administration of PGE2 together with LPS results in a reduction in the levels of CCL3 and CCL4 released in the peritoneal fluid, a reduction in the number of dendritic cells accumulating in the peritoneal cavity, and a reduction in CCL3 amount per cell in the peritoneal cell population. These results suggest that one of the mechanisms by which endogenous PGE2 acts as an anti-inflammatory agent, is the inhibition of inflammatory chemokine release from activated dendritic cells, preventing the excess accumulation of activated immune cells.

Key Words: lipid mediators • inflammation • bone marrow-derived dendritic cells • chemokines


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In the last decade, dendritic cells (DCs) emerged as major antigen presenting cells capable of activating naïve T cells and bridge innate and adaptive immunity [1 , 2 ]. Following exposure to pathogens or in response to inflammatory cytokines, DCs undergo maturation. Mature DCs produce a variety of cytokines and chemokines [3 4 5 ] and become efficient antigen-presenting cells [6 7 8 9 10 11 12 ]. DCs utilize a relatively small number of Toll-like receptors (TLRs) to recognize different pathogen-associated molecular patterns (PAMPs) [reviewed in 13 ], and respond with specific gene expression programs, in addition to the common core response [reviewed in 14 , 15 ]. Cytokines such as TNF{alpha}, IL-1ß, and IL-6, and chemokines such as CCL3 (MIP-1{alpha}), and CCL4 (MIP-1ß) are part of the common core response. Activated DCs produce CCL3 and CCL4 early, and at high levels [5 ], and CCL3/CCL4 act as chemotactic factors for cells expressing CCR1, 2, 4, and 5 receptors. Inflammatory immune cells such as monocytes/macrophages, DCs, activated Th1 and Th2 cells, as well as NK cells and eosinophils, express one or more of these receptors, and respond by accumulating at the infection site.

Although an inflammatory response is necessary for the containment and elimination of pathogens, uncontrolled inflammation is highly detrimental to the host, leading to a variety of pathological conditions. Several endogenous anti-inflammatory agents contribute to immune homeostasis, by eliminating or deactivating excess inflammatory cells. Cytokines such as IL-10, IL-13, and TGF-ß [reviewed in 16 ], neuroendocrine factors such as glucocorticoids and neuropeptides [reviewed in 17 , 18 ], and lipid mediators such as prostaglandins [reviewed in 19 , 20 ], function as endogenous anti-inflammatory agents, causing apoptosis of activated T cells, preferential differentiation into Th2 as opposed to Th1 effectors, and/or deactivation of macrophages, microglia, and dendritic cells.

PGE2, generated from arachidonic acid by cyclo-oxygenases (Cox) and prostaglandin E synthase [21 ], affects both macrophage and T cell activation, exerting a general anti-inflammatory effect [reviewed in 20]. PGE2 has different effects on immature vs. mature DCs. In immature DCs, PGE2 cooperates with inflammatory cytokines such as TNF{alpha}, IL-1ß, and IL-6 to promote DC maturation [22 , 23 ]. In contrast, most of the PGE2 effects on mature DCs are anti-inflammatory. PGE2 inhibits the production of the pro-inflammatory cytokine IL-12p70 and promotes the release of the anti-inflammatory cytokine IL-10 from stimulated DCs, skewing CD4 T cell differentiation toward Th2 effectors [24 25 26 27 ]. In agreement with the proposed Th2 bias, PGE2 inhibits the production of CXCL10 (IP-10), a Th1 chemoattractant and promotes CCL22 (MDC), a preferential Th2 chemoattractant, in splenic DCs and macrophages [28 ].

PGE2 has been reported to inhibit the production of several pro-inflammatory CKs in macrophages and astrocytes [29 , 30 ]. However, to our knowledge, this is the first report of the inhibitory effect of PGE2 on CCL3 and CCL4 release from activated DCs. We examined the effects of PGE2 on the CCL3/CCL4 production by bone marrow-derived murine DCs stimulated through TLR4 (lipopolysaccharide; LPS) or TLR2 (peptidoglycan; PGN). Our results indicate that PGE2 inhibits CCL3 and CCL4 production, in a dose- and time-dependent manner, both at mRNA and protein level. Agonist studies identified EP2 and EP4 receptors as mediators of the inhibitory effect of PGE2, and purified DCs express these receptors both at the mRNA and protein levels. The in vitro inhibitory effect of PGE2 was confirmed in vivo, where i.p. administration of PGE2 together with LPS resulted in a reduced accumulation of DCs in the peritoneal cavity and a lower level of CCL3 expression both in the peritoneal fluid and peritoneal exudate cells.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Mice
Male B10.A mice 6-8 weeks old were purchased from Jackson Laboratories (Bar Harbor, ME) and were maintained in the Rutgers State University (Newark, NJ) animal facility under pathogen free conditions. The animals used in the in vivo experiments were 22-23 g in weight.

Reagents
LPS (Escherichia coli O26:56), peptidoglycan (PGN from Staphylococcous aureus), prostaglandin E2 (PGE2), and avidin-peroxidase were purchased from Sigma (St. Louis Chemical Company, MO). TMB Elisa substrate was purchased from Pierce (Rockford, IL). Capture and biotinylated anti-mouse MIP-1ß antibody were purchased from PharMingen (San Diego, CA). Recombinant murine MIP-1ß was purchased from R&D systems (Minneapolis, MN). Purified and biotinylated anti-mouse MIP-1{alpha} antibodies and recombinant mouse MIP-1{alpha} were purchased from Peprotech (Rock Hill, NJ). The EP receptor agonists butaprost, misoprostol, sulprostone, ibuprofen (a nonspecific Cox inhibitor) and NS-398 (a specific Cox 2 inhibitor) were purchased from Cayman Chemical (Ann Arbor, MI). The J558L plasmacytoma cell line transfected with the murine GM-CSF was kindly provided by Dr. Jeffrey Ravetch (Rockefeller University, New York). The following antibodies were used for FACS analysis: PE-conjugated anti-CD11c (PharMingen), rabbit anti-mouse MIP-1{alpha} (Peprotech), rabbit anti-murine EP2 and EP4 polyclonal antibodies (Cayman Chemicals, Ann Arbor, MI), and FITC-conjugated goat anti-rabbit IgG (Sigma) as indirect reagent.

Generation and purification of DCs from bone marrow
DCs were generated in vitro from B10.A mouse bone marrow according to Lutz et al. [31 ] with slight modifications. Briefly, femura and tibiae were removed from 6-8 weeks old male B10.A. Both ends of the bone were cut open, and bone-marrow cells were flushed out and washed with ice-cold RPMI 1640 medium (Gibco-BRL, Grand Island, NY). BM cells (2 x 106 cells) were cultured in 100 mm petri dishes containing 10 ml RPMI 1640 medium supplemented with 10% heat-inactivated FBS (Atlanta Biologicals, Norcross, GA), 2 mM L-glutamine, 50 µM 2-mercaptoethanol, and supernatants from murine GM-CSF transfected J558L cells. The final concentration of GM-CSF in the DC complete medium is 20 ng/ml. After three days, another 10 ml of complete medium containing GM-CSF was added to each dish. At day 8 the nonadherent cells were harvested and used as immature DCs in some experiments. The percentage of CD11c+ DCs in the nonadherent population averaged 70% by FACS analysis. To ensure the purity of DCs generated from bone morrow described above, CD11c+ DCs were purified by immunomagnetic sorting using anti-CD11c-coated magnetic beads and the autoMACS system, according to the manufacturer’s instructions (Miltenyi Biotech, Bergish-Gladbach, Germany). The purity of the sorted cells was determined by FACS analysis (>96% for CD11c+ cells).

MIP-1{alpha}, MIP-1ß, and endogenous PGE2 measurements by ELISA
Purified DCs were cultured in 24-well culture plates at 1 x 105 cells per well in a final volume of 1 ml and stimulated with different concentrations of LPS (0-10 µg/ml) or PGN (0-50 µg/ml) in the presence or absence of PGE2 (10-6-10-10 M). Cell-free supernatants were harvested at the designated time points and subjected to MIP-1{alpha} and MIP-1ß ELISA assays. For the detection of MIP-1{alpha}, 0.5 µg/ml purified polyclonal antibody was used to coat plates, followed by detection with the biotinylated anti-mouse MIP-1{alpha} Ab (0.2 µg/ml). MIP-1ß was assayed with AG5-2 as capture Ab (4 µg/ml) followed by detection with the biotinylated anti-mouse MIP-1ß Ab (2 µg/ml). The sensitivity of ELISAs for MIP-1{alpha} and MIP-1ß is 20 pg/ml. PGE2 production was measured by ELISA (Cayman Chemicals), as recommended by the manufacturer. The detection limit for the PGE2 ELISA is 15 pg/ml.

Stimulation of immature DCs and RNA preparation
In some experiments, purified CD11c+ DCs were cultured in petri dishes at a concentration of 1 x 106 cells/ml and stimulated with LPS or PGN in the presence or absence of PGE2 for 3 h. Cells were collected, and total RNA was isolated with the Ultraspec RNA reagent (Biotecx Laboratories, Houston, TX), as recommended by the manufacturer. In other experiments, unpurified nonadherent cells from BM cultures (day 8) were transferred to new petri dishes (1 x 106 cells/ml) and treated as above. Three hours later, cells were harvested, followed immediately by purification of CD11c+ DCs and RNA extraction.

RNase protection assay (RPA)
RNase protection assay (RPA) was performed with 3-5 µg of total RNA using the mCK-5 multiprobe template and the Riboquant MultiProbe RPA system (BD PharMingen), following the manufacturer’s instructions. In brief, [{alpha}-32P] UTP-labeled antisense RNA probes were synthesized by using the mCK-5 multiprobe in vitro transcription system. Antisense RNA probes were purified by phenol/chloroform extraction and hybridized with the RNA samples at 56°C overnight. Unhybridized single-stranded RNA was digested by RNase treatment. The undigested RNAs were separated on a 5% denaturing polyacrylamide gel. The gel was dried, and the intensity of the radioactive bands was quantitated using a phosphoimager: SI (Molecular Dynamics, Sunnyvale, CA). Data are expressed as a ratio of MIP-1{alpha} (or MIP-1ß) mRNA to the housekeeping gene GAPDH mRNA from the same sample to correct for any error in RNA loading.

Real-time PCR
The SYBR green-based real-time PCR technique was used to detect the expression of chemokines or prostanoid receptors (EPs) in DCs. Cells were treated as described above, and RNA was prepared from purified CD11c+ DCs. One µg of total RNA was reverse transcribed into cDNA in the presence of 200 units of MMLV-RT, 40 units of RNasin, 1 µg of random primers, 0.5 mM dNTPs, 3 µg of BSA, and 1 x MMLV reaction buffer (Promega, Madison, WI) in a total volume of 30 µl at 42°C for 1 h. cDNA was then diluted 10 times for real-time PCR. The PCR mixture consists of 2 µl diluted cDNA, 5 µl SYBR green-containing PCR master mixture (2x) and 150 µM of each primer in a total volume of 10 µl. The specific primers for real-time PCR were designed by using the Primer ExpressTM software from Applied Biosystems, Inc. (Foster City, CA), and are as follows: MIP-1{alpha} sense 5'-ACTGCCTGCTGCTTCTCCTACA-3' and antisense 5'- AGGAAAATGACACCTGGCTGG-3'; MIP-1ß sense 5'-AAACCTAACCCCGAGCAACA-3' and antisense 5'-CCATTGGTGCTGAGAACCCT- 3'; EP1 sense 5'-TTAACCTGAGCCTAGGGGATG-3' and antisense 5'-CGCTGGTGATGTGCCATTATC-3'; EP2 sense 5'-TGCAAGAGTCGTCAGTGGCT-3' and antisense 5'-AACAGTGCCAGTGCGATGAG-3'; EP3 sense 5'-CAATCAGATGTCGGTTGAGCA-3' and antisense 5'-AGATCTGGTTCAGCGAAGCC-3'; EP4 sense 5'-TTTCTTCGGTCTGTCGGGTC-3' and antisense 5'-CGCTTGTCCACGTAGTGGCT-3'. Real-time PCR was performed using the ABI PRISM 7900HT sequence detection system (Applied Biosystems), and the cycling conditions used were 95°C for 15 s, 60°C for 1 min for 40 cycles, followed by a melting point determination or dissociation curve that results in a single peak if the amplification is specific. The expression level for each gene is showed by the cycle numbers needed for the cDNA to be amplified to reach a threshold. At the same time, the cycle numbers are transferred into arbitrary amounts of DNA using a standard curve generated for each pair of primers, and the results are normalized to the housekeeping gene GAPDH mRNA from the same sample.

In vivo experiments
B10.A male mice (3 mice/group) were injected with a single i.p. dose of LPS (25 µg/mouse; 1.1 mg/kg) in the presence or absence of different amounts of PGE2 (0.5-50 nmoles/mouse; 22 nmoles-2.2 µmoles/kg). Six hours later, the mice were killed and peritoneal exudates were obtained by lavage with 2 ml of ice-cold PBS containing 3mM EDTA. Cell-free supernatants were harvested and assayed for MIP-1{alpha} and MIP-1ß production by ELISA, as described above. Control mice received PBS or PGE2 without LPS. Peritoneal exudate cells obtained 5 h following inoculation were subjected to FACS analysis for surface CD11c expression and intracellular MIP-1{alpha} production. For the determination of PGE2 in the peritoneal fluid, groups of three mice were injected with medium or LPS (25 µg/animal), and peritoneal fluid was obtained following lavage with 1 ml of ice-cold PBS containing 3 mM EDTA at various time points. The cell-free supernatants were assayed for PGE2 by ELISA.

FACS analysis
Cells were subjected to analysis by using a 3-color FACSCalibur (BD Biosciences, Mountain View, CA) after staining with anti-CD11c-PE and rabbit anti-mouse MIP-1{alpha} followed by goat anti-rabbit IgG-FITC. The specificity of the primary Abs was established with appropriate isotype-matched controls. Data were collected for 10,000 cells and analyzed using Cellquest software from BD Biosciences.

Statistical analysis
Results are expressed as the mean ± SEM. Analysis of data was performed using the Student’s t test with p < 0.05 as the minimum significant level.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
PGE2 inhibits LPS-induced MIP-1{alpha} and MIP-1ß production
To investigate the effects of PGE2 on MIP-1{alpha} and MIP-1ß production, purified CD11c+ DCs were stimulated with different concentration of LPS (0-10 µg/ml) in the presence or absence of various doses of PGE2 and the amounts of CKs released in the culture supernatants were assayed by ELISA at different time points. PGE2 inhibited both MIP-1{alpha} and MIP-1ß production in a dose- and time-dependent manner. The inhibitory effect is exerted over a large LPS concentration range (from 0.01 to 10 µg/ml) (Fig. 1A 1B ). The inhibitory effect was dose-dependent, with significant inhibition for 10-6-10-8M PGE2 (Fig. 1C 1D) . The time-response curves indicate that MIP-1{alpha} and MIP-1ß release was inhibited by PGE2 as early as 3 h, with a maximum inhibitory effect after 6 h of culture (Fig. 1E 1F) . The inhibitory effect of PGE2 was more pronounced for MIP-1{alpha} than MIP-1ß, at all time points and for all PGE2 concentrations.



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  		Figure 1. PGE2 inhibits MIP-1{alpha} and MIP-1ß production from LPS-stimulated dendritic cells (DCs). Purified, CD11c+ bone marrow-derived DCs were stimulated with different concentrations of LPS (0-0 µg/ml) in the presence or absence of PGE2 (10-7 M). Supernatants were harvested 18 h later and subjected to ELISA for (A) MIP-1{alpha} or (B) MIP-1ß. DCs were treated with different concentrations of PGE2 (10-6–10-10 M) in the presence of LPS (1µg/ml). Supernatants were collected 18 h later and subjected to ELISA for (C) MIP-1{alpha} or (D) MIP-1ß. DCs were treated with PGE2 (10-7 M) and LPS (1µg/ml). Supernatants were collected at different time points [3, 6, 9, 24, 48 h] and assayed for (E) MIP-1{alpha} production or (F) MIP-1ß. * indicates statistically significant differences compared with the LPS-treated group (p<0.05).

 
PGE2 inhibits PGN-induced MIP-1{alpha} and MIP-1ß production
Because LPS and PGN stimulate DCs through different TLR, we investigated next whether PGE2 inhibits MIP-1{alpha} and MIP-1ß release following PGN stimulation. Purified CD11c+ DCs were stimulated with different concentration of PGN (0.1-50 µg/ml) in the presence or absence of different concentration of PGE2 (10-6–10-10 M). Similar to the results obtained for LPS, PGE2 inhibits PGN induced MIP-1{alpha} and MIP-1ß (Fig. 2 ). The inhibitory effect is exerted over a large PGN concentration range (from 0.1 to 50 µg/ml) (Fig. 2A 2B) , and at all time points (Fig. 2E 2F) . The inhibition is dose-dependent, with significant values for 10-6 and 10-7 M PGE2 (Fig. 2C 2D) .



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  		Figure 2. PGE2 inhibits MIP-1{alpha} and MIP-1ß production from PGN-stimulated dendritic cells (DCs). DCs were stimulated with different concentrations of PGN (0-50 µg/ml) in the presence or absence of PGE2 (10-7 M). Supernatants were harvested 18 h later and subjected to ELISA for (A) MIP-1{alpha} or (B) MIP-1ß. DCs were treated with different concentrations of PGE2 (10-6–10-10 M) and PGN (10µg/ml). Supernatants were collected 18 h and subjected to ELISA for (C) MIP-1{alpha} or (D) MIP-1ß. DCs were treated with PGE2 (10-7 M) and PGN (10 µg/ml). Supernatants were collected at different time points and tested for (E) MIP-1{alpha} or (F) MIP-1ß. * indicates statistically significant differences compared with the LPS-treated group (p<0.05).

 
The effects of the purification process on DCs
Because we observed high basal levels of MIP-1{alpha} and MIP-1ß in medium controls of purified DCs, we investigated whether the purification process with anti-CD11c microbeads might activate immature DCs. Immature DCs treated in two different ways were compared in terms of CK expression. The first treatment consisted of nonpurified DCs stimulated with LPS for 3 h, followed by purification of CD11c+ cells immediately before RNA extraction (Group I). In the second treatment, CD11c+ cells were first purified, followed by LPS treatment for 3 h and RNA extraction (Group II). Three micrograms total RNA was subjected to RPA analysis. There was a higher expression of chemokines in the medium control in Group II (purified before the LPS treatment) compared with Group I (purified after LPS treatment), particularly for MIP-1ß and IP-10 (Fig. 3A ). MIP-1{alpha} and -ß expression from Fig. 3A were quantified in Fig. 3B . These results were confirmed by real-time PCR (Fig. 3C) . Although PGE2 exerts an inhibitory effect on MIP-1{alpha} and -1ß expression, the inhibition is more pronounced for cells purified after the LPS treatment (Group I), where CK levels are much lower in medium controls (Fig. 3B 3C) . Because the purification process appears to induce the expression of some CK genes, in experiments related to MIP-1{alpha}/1ß expression at the RNA level, we treated nonpurified, nonadhenrent immature DCs (70% CD11c+ on average) with LPS for 3 h, followed by separation of CD11c+ cells (>96% CD11c+), preparation of total RNA, and real-time PCR analysis.



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  		Figure 3. The effects of purification with anti-CD11c+ magnetic beads on CK expression. (A) Immature, nonadherent cells (70% CD11c+) were treated with LPS (1µg/ml) +/- PGE2 for 3 h, followed by purification of DCs with CD11c+ magnetic beads, and total RNA extraction (Group I). Immature DCs were purified with anti-CD11c+ magnetic beads (>96% CD11c+ by FACS), and treated with LPS (1µg/ml) in the presence or absence of PGE2 (10-7M). RNA was extracted 3 h later and subjected to RPA (Group II). One representative experiment out of two is shown. (B) Quantitative analysis of MIP-1{alpha} and (C) ß mRNA expression from Fig. 3A. Analysis of MIP-1{alpha} and -ß expression in the same samples from Fig. 3A by real-time PCR. One representative experiment out of four is shown.

 
PGE2 inhibits MIP-1{alpha} and MIP-1ß mRNA expression
Nonpurified nonadherent immature DCs were treated with different concentration of LPS in the presence or absence of PGE2. Three hours later, the CD11c+ cells were isolated for RNA extraction. The expression of MIP-1{alpha} and MIP-1ß was determined by real-time PCR. PGE2 inhibits expression of both CKs over a range of LPS doses (Fig. 4A 4B ), and the inhibition is dose-dependent (Fig. 4C 4D) . We then proceeded to investigate at what time point PGE2 exerts its inhibitory effect. DCs were treated with LPS, and PGE2 was added at different times. The inhibition was observed when PGE2 was added simultaneously with LPS or when the cells were pretreated with PGE2 for 30 min. In contrast, there was no inhibitory effect when PGE2 was added 30 min after LPS (Fig. 4E 4F) .



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  		Figure 4. PGE2 inhibits MIP-1{alpha} and MIP-1ß mRNA expression. Immature DCs were treated with different concentrations of LPS (0-10 µg/ml) in the presence or absence of PGE2 (10-7 M). Three hours later, the CD11c+ cells were isolated, and RNA was extracted and assayed by (A) real-time PCR for MIP-1{alpha} or (B) MIP-1ß expression. One representative experiment out of four is shown. (C)-(D) DCs were treated with LPS (1µg/ml) and different concentration of PGE2 (10-6–10-10 M) for 3 h. RNA was assayed by real-time PCR. One representative experiment out of four is shown. (E)-(F) DCs were pretreated with PGE2 (10-7 M) for 30 min before LPS (1 µg/ml) (before), exposed to LPS and PGE2 at the same time (same), or treated with PGE2 30 min after LPS (after). CD11c+ cells were purified 3 h later, and RNA was extracted and assayed for MIP-1{alpha} and MIP-1ß expression.

 
The PGE2 inhibition of MIP-1{alpha} and MIP-1ß is mediated by EP2, and possibly EP4 receptors
Several PGE2 receptors (EP1, 2, 3, and 4) are expressed in various tissues. To determine which of the EP receptors are involved in the inhibitory effect of PGE2 on MIP-1{alpha}/MIP-1ß expression, we used receptor agonists. Sulprostone, a relatively specific agonist for EP1 (Ki=21 nM) and EP3 (Ki=0.6 nM), did not inhibit LPS induced MIP-1{alpha} and MIP-1ß expression, even at concentrations up to 10-6M (Fig. 5A 5B ). On the other hand, butaprost, a selective agonist for the EP2, inhibits MIP-1{alpha} and MIP-1ß expression, similar to PGE2 (Fig. 5A 5B) . Misoprostol binds EP3 and EP4 receptors at low concentrations (Ki =67 nM), and also EP1 and EP2 receptors at higher concentrations (with Ki =120, 250 nM for EP1 and EP2, respectively). We treated DCs with different concentrations of misoprostol (50 nM to 1 µM). Misoprostol inhibited MIP-1{alpha}/MIP-1ß expression in a dose-dependent manner (Fig. 5A 5B) . These findings suggest that EP2 and EP4 receptors mediate the inhibitory effect of PGE2 on MIP-1{alpha}/MIP-1ß.



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  		Figure 5. PGE2 inhibition of MIP-1{alpha} and MIP-1ß expression is mediated through EP2 and EP4 receptors. (A) Nonadherent, immature DCs were stimulated with LPS (0.1 µg/ml) in the presence or absence of 0.1µM PGE2, 1 µM sulprostol (agonist for EP1/EP3), 10 µM butaprost (an EP2 agonist), and different concentrations of misoprostone (1:1000nM; 2:200nM; 3:50nM). CD11c+ cells were isolated 3 h later, RNA was prepared and subjected to real-time PCR for MIP-1{alpha} and MIP-1ß. (B) Cells were treated the same way as in Fig. 6A , except that PGN (1µg/ml) was used instead of LPS. For both (A) and (B), one representative experiment out of three is shown.

 
There are no reports at the present time regarding the expression of PGE2 receptors in bone marrow-derived murine DCs. By using real-time PCR, we observed expression of EP1, EP2 and EP4 in immature DCs, with the cycle Ct of 24, 26, and 25 for EP1, EP2, and EP4, respectively (Fig. 6A ). In contrast, the cycle Ct for EP3 is 29, suggesting that EP3 is probably not expressed, or expressed at very low levels in immature murine DCs. The presence of EP2 and EP4 on immature DCs was confirmed by FACS analysis (Fig. 6B) .



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  		Figure 6. Expression of EP receptors in immature DCs. (A) Total RNA prepared from purified immature DC was subjected to real-time PCR with primers specific for EP1, 2, 3, and 4 as described in Materials and Methods. Results are shown as amplification plots for each receptor. One representative experiment out of two is shown. (B) Immature DCs were harvested and subjected to FACS analysis for EP2 and EP4 receptors. Cells were stained for both CD11c and EP2/EP4. Cells were first gated for CD11c marker and then analyzed for expression of EP2 and EP4. The dotted line represents the isotype controls. One representative experiment out of two is shown.

 
Endogenous PGE2 does not affect CCL3 and CCL4 production
To determine whether endogenous PGE2 produced in response to LPS plays a role in the inhibition of CCL3/4 production, we measured the amounts of PGE2 released by BM-DCs following LPS stimulation. Endogenous PGE2 levels were significantly upregulated as early as 6 h, reaching a plateau between 12 and 24 h (Fig. 7A ). However, exogenous PGE2 added 30 min after LPS did not inhibit CCL3/4 production (Fig. 4E and 4F) . This suggests that endogenous PGE2 might not play a role in the in vitro BM-DCs cultures, because of insufficient release within minutes after LPS stimulation. This was confirmed by treatment with LPS in the presence of different concentrations of ibuprofen (a Cox 1/2 inhibitor) and NS-398 (a specific Cox 2 inhibitor). Neither ibuprofen, nor NS-398 affected the levels of CCL3 and CCL4 released in response to LPS (Fig. 7B) .



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  		Figure 7. Endogenous PGE2 does not affect MIP-1{alpha} and MIP-1ß production. (A) Purified DCs (0.5 x 106/ml) were stimulated with LPS (1 µg/ml) and supernatants were harvested at different time points (0, 3, 6, 12, 24, 48 h). Unstimulated DCs were used as a control. PGE2 quantification in the supernatants was carried out using ELISA. (B) Purified DCs (0.5 x 106 cells/ml) were stimulated with LPS (1 µg/ml) in the presence of different concentrations of ibuprofen (10-5 to 10-8M) or NS398 (10-5 to 10-9M). Supernatants were collected after 18 h and subjected to MIP-1{alpha} and MIP-1ß ELISAs. One experiment out of two is shown.

 
In vivo effects of PGE2 on MIP-1{alpha} and MIP-1ß production
To assess whether PGE2 affects the in vivo production of chemokines, we inoculated B10.A mice i.p. with LPS (25 µg/animal; 0.1 ml) and PGE2 (0.5-50 nmoles/animal). Peritoneal fluid was harvested and subjected to ELISA for MIP-1{alpha} and MIP-1ß. PGE2 inhibits the in vivo production of MIP-1{alpha} and MIP-1ß (Fig. 8A 8B ). In addition, peritoneal cells harvested from mice injected with both LPS and PGE2 express less intracellular MIP-1{alpha} on a per cell basis compared with LPS alone (Fig. 8D) .



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  		Figure 8. In vivoeffects of PGE2 on LPS-induced MIP-1{alpha} and MIP-1ß production. B10.A mice were injected i.p. with LPS (25 µg), or LPS plus different concentrations of PGE2 (0.5-50 nmoles/mouse). Six hours later, peritoneal exudate fluid was obtained and assayed (A) for MIP-1{alpha} and (B) MIP-1ß by ELISAs. One representative experiment out of two is shown. (C) Mice were injected i.p. with LPS (25 µg), LPS plus PGE2 (5 nmoles/mouse) or PBS. Five hours later, peritoneal exudate cells were obtained, stained with PE-labeled anti-CD11c Ab and subjected to FACS analysis (10,000 cells). Control (time 0)-mice were injected with LPS and killed immediately. PBS (5 h)-mice were injected with LPS and killed 5 h later. (D) Mice were injected i.p. with LPS (25 µg), or LPS plus PGE2 (5 nmoles/mouse). Five hours later, peritoneal exudate cells were obtained, permeabilized, stained for intracellular MIP-1{alpha}, and analyzed by FACS. The average MCF (mean channel fluorescence) for MIP-1{alpha} staining is shown. * indicates statistically significant differences compared with the LPS treated group (p<0.05). One representative experiment out of three is shown. (E) Mice were injected i.p. with LPS (25 µg/animal). Peritoneal fluid (1 ml) was obtained at various time points and assayed for PGE2 by ELISA. Time 0 represents values obtained from animals injected with medium or not injected. One representative experiment out of two is shown.

 
To address the question whether PGE2 affects DC accumulation in the peritoneal cavity, B10.A mice were inoculated i.p. with LPS or LPS plus PGE2, and the numbers of CD11c+ cells in the peritoneal exudate were determined by FACS. Although CD11c+ cells are relatively rare in the peritoneal exudate population, LPS induces an increase in DCs (at 5 h) compared with time 0 and compared with controls inoculated with PBS. In contrast, administration of PGE2 results in a significant decrease in the number of CD11c+ cells (Fig. 8C) .

The in vivo administration of LPS leads to endogenous PGE2 production, and therefore, both endogenous and exogenous PGE2 could contribute to the effects described above. To determine the levels and time course for the generation of endogenous PGE2, we treated groups of B.10 mice with LPS, and harvested peritoneal fluid at various time points. Controls consisted of mice injected with medium, or not injected at all. Both medium-injected and noninjected mice averaged 0.8 ng/ml of PGE2. There was little increase if any, up to 2 h following LPS administration. An increase in endogenous PGE2 was observed 4 h after LPS administration (Fig. 8E) . However, even at 4 h, the endogenous PGE2 concentration was 50-fold lower than the minimum dose of exogenous PGE2 used in our experiments (Fig. 8A and 8B) .


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Bridging innate and adaptive immunity, DCs eliminate pathogens, attract various immune cells to the site of infection, and following maturation, migrate to T cell areas where they present antigen and activate naïve T cells. Production of proinflammatory chemokines such as CCL3, CCL4, and CCL5 results in the attraction of activated T cells, monocytes/macrophages, NK cells, and eosinophils. Although necessary for the containment of pathogens, uncontrolled production of proinflammatory chemokines and cytokines is detrimental to the host. Together with other endogenous deactivators such as antiinflammatory cytokines, and neuroendocrine factors, PGE2 plays an important role in controlling inflammation.

PGE2 was reported to control CCL3 (MIP-1{alpha}), CCL4 (MIP-1ß), and CCL5 (RANTES) production in human astrocytes and macrophages [29 , 30 ]. In this study, we report on the PGE2 inhibition of CCL3 and CCL4 production in murine bone marrow-derived DCs. The inhibition was exerted in a dose-dependent manner at both mRNA and protein level. The effect on MIP-1{alpha} and MIP-1ß appears to be specific. However, although PGE2 did not affect MIP-2, IP-10, MCP-1 and RANTES expression in our experimental system (Fig. 3A) , this has to be confirmed at several time points.

Four heptahelical transmembrane receptors designated EP1-4, coupled to different G proteins, mediate PGE2 signaling [reviewed in 32 ]. The receptors have unique expression patterns, and because of coupling to different G proteins, use different transduction pathways. EP2 and EP4, both cAMP inducers, have been previously reported to mediate the effects of PGE2 in immune cells [30 , 33 34 35 ]. We approached the question of the receptors involved in the inhibitory effect of PGE2 by using receptor agonists. Both butaprost (a specific EP2 agonist), and misoprostol (an EP3 and EP4 agonist at low concentrations) mimicked the inhibitory effect of PGE2. On the other hand, sulprostone (an EP1 and EP3 agonist) did not inhibit CCL3 and CCL4 production. These results suggest that EP2 and EP4 mediate the inhibitory effects of PGE2 on DCs. Real-time PCR and FACS analysis confirmed the expression of EP2 and EP4 in immature DCs at the time of PGE2 treatment. The relative contribution of the two receptors to the effect of PGE2 on CCL3 and CCL4 production in DCs will have to be determined using DCs derived from EP2 and EP4 knockout mice.

Our experiments confirm that PGE2 inhibits CCL3 and CCL4 at both the mRNA and protein levels. Interestingly, the levels of CCL3 and CCL3 mRNA expression in DCs increased following purification through immunomagnetic methods. This might be due to partial activation upon ligation of the CD11c molecule. Indeed, the CD11c integrin has been proposed to be part of the LPS activation cluster [36 ], and therefore, ligand binding to CD11c might induce the expression of certain genes. It should be noted however, that CD11c ligation does not induce TNF{alpha} expression in DCs (unpublished results). To minimize CCL3 and CCL4 expression in controls, we stimulated bone marrow-derived nonadherent, unpurified day 8 cells (70% CD11c+ on average) with LPS or PGN in the absence or presence of PGE2 for 3 h, followed by purification of CD11c+ DCs and RNA extraction. Although the RNA data originate only from the CD11c+ DCs, it is theoretically possible that the effects of PGE2 were mediated through the remaining 30% CD11c- cells. However, this is improbable, since for the CCL3 and CCL4 protein determinations, we purified the CD11c+ DCs before exposure to LPS/PGN and PGE2, and the results were similar to those observed in the real-time PCR experiments.

Stimulation of DCs through different TLR receptors results in the expression of an overlapping group of genes including CCL3, CCL4, TNF{alpha}, and IL-6 through the MyD88/IRAK/NFkB-P38-JNK pathways [15 , 37 ]. EP2 and EP4 are linked to cAMP induction, and subsequent PKA activation. The relationship between cAMP-inducing receptors and effects on NFkB, P38, or JNK is not clear. We have shown previously that another endogenous cAMP inducing agent, the neuropeptide VIP, inhibits expression of proinflammatory chemokines, including CCL3 and CCL4, in LPS-stimulated macrophages and microglia and that these effects correlate with an inhibition of NFkB binding [38 , 39 ]. In addition to PKA activation, other intracellular pathways might be involved. A recent report suggests that whereas EP2 activates the T cell factor (Tcf) transcriptional factor through PKA, EP4 signals mainly through the phosphatidylinositol 3-kinase (PI3K) pathway [40 ]. Whether this applies to the effect of PGE2 on CCL3 and CCL4 expression in LPS/PGN-stimulated DCs remains to be established.

LPS stimulation has been reported to induce endogenous PGE2 production in DCs [41 ]. Therefore, the contribution of endogenous PGE2 cannot be ignored. However, our results indicate that exogenous PGE2 added 30 min after LPS does not inhibit CK production, and unpurified or purified CD11c+ DCs do not secrete significant levels of endogenous PGE2 immediately following LPS stimulation. In addition, DCs stimulated with LPS in the presence or absence of Cox inhibitors produced the same amounts of CCL3 and CCL4. Similar conclusions were reached by Jozefowski et al. [42 ] for the production of cytokines by bone marrow-derived DCs.

The biological significance of the PGE2-induced reduction in CCL3 and CCL4 expression is underlined by the in vivo experiments. Administration of exogenous PGE2 reduced the accumulation of DCs in the peritoneal cavity and the amounts of CCL3 and CCL4 released in the peritoneal fluid. PGE2 also reduced the amounts of CCL3 in the peritoneal exudate cells on a per cell basis. For the in vivo experiments, the role of endogenous PGE2 generated following LPS administration cannot be discounted. We investigated the accumulation of DCs and CCL3/CCL4 production in the peritoneal cavity at 5 and 6 h, respectively, whereas an increase in endogenous PGE2 levels (still 50-fold lower than the dose of exogenous PGE2) was apparent 4 h after LPS administration. These results confirm an anti-inflammatory in vivo effect of PGE2, and support the hypothesis that endogenous PGE2, released at later time points during an inflammatory reaction, acts as a natural anti-inflammatory agent, preventing the further amplification of the inflammatory response.


   ACKNOWLEDGEMENTS
 
This work was supported by grants AI47325 and AI052306 (DG), and by the Johnson&Johnson Fellowship for Neuroimmunology 2002-2003 (HJ and EV).

Received March 24, 2003; revised July 8, 2003; accepted July 9, 2003.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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