The JI PBL Intereron Source
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     
 

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Baratelli, F. E.
Right arrow Articles by Dubinett, S. M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Baratelli, F. E.
Right arrow Articles by Dubinett, S. M.
The Journal of Immunology, 2004, 173: 5458-5466.
Copyright © 2004 by The American Association of Immunologists

Prostaglandin E2-Dependent Enhancement of Tissue Inhibitors of Metalloproteinases-1 Production Limits Dendritic Cell Migration through Extracellular Matrix1

Felicita E. Baratelli2, Nathalie Heuzé-Vourc’h2, Kostyantyn Krysan, Mariam Dohadwala, Karen Riedl, Sherven Sharma and Steven M. Dubinett3

Lung Cancer Research Program, Jonsson Comprehensive Cancer Center, and the Division of Pulmonary and Critical Care Medicine, Department of Medicine, David Geffen School of Medicine, University of California, Los Angeles, CA 90095


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Dendritic cell (DC) migration is crucial for the initiation of immune responses. The balance between metalloproteinases (MMP) and tissue inhibitors of metalloproteinases (TIMP) has been shown to modulate DC migration. PGE2, which is overproduced in a wide variety of human malignancies, has been implicated in MMP and TIMP regulation in various cells, including monocytes. In the present study, we hypothesized that tumor-derived PGE2 would affect DC migratory capacity through the extracellular matrix (ECM) by altering MMP and TIMP balance. Treatment of monocyte-derived immature DC with exogenous PGE2 induced TIMP-1 secretion but not MMP-9 production and was correlated with reduced DC migration through ECM. Because recombinant TIMP-1 replicated PGE2 inhibition of DC migration while anti-TIMP-1 neutralizing Ab reversed it, we conclude that PGE2-mediated induction of TIMP-1 was responsible for the reduced migration of PGE2-treated DC. Similarly, DC cultured for 48 h in supernatants from cyclooxygenase-2 overexpressing lung cancer cells that secrete high levels of PGE2, exhibited decreased migration through ECM. Finally, analysis of E prostanoid receptor expression and their selective inhibition revealed that the enhanced TIMP-1 secretion in PGE2-treated DC was mediated predominantly by the E prostanoid receptor 2. These findings indicate that PGE2-dependent enhancement of TIMP-1 production causes reduced migration of DC through ECM.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Dendritic cells (DCs)4 act as sentinels of the immune system initiating and controlling adaptive immune responses (1, 2). In view of their crucial role, it is important to understand how DCs patrol in the periphery and to define mechanisms that control and regulate DC trafficking. DC migration is a complex process that involves a coordinate activation of different classes of effector molecules: adhesion molecules, chemotactic factors, and proteases (2, 3, 4). Among chemotactic factors, chemokines have been the focus of extensive investigations as they serve as an important signal for the regulation of DC migration in vitro and in vivo. The expression and response to chemokines vary depending on DC origin and maturation status (5, 6).

Proteases are also important in orchestrating cell migration as they facilitate cell movement through the extracellular matrix (ECM). Among them, matrix metalloproteinases (MMP) have been identified as key secreted enzymes for the degradation of ECM components and are considered physiologic mediators of both normal and tumor cell migration (7, 8, 9, 10). In particular, the gelatinases MMP-2 and MMP-9 cleave type IV collagen, which constitutes a backbone for attachment of other basement membrane components (10, 11). Because MMPs have the potential to cause deleterious tissue destruction, their enzymatic activity is tightly controlled at different levels: gene transcription, activation of the zymogen form, and inhibition by broad-spectrum inhibitors ({alpha}2-macroglobulin) or the specific tissue inhibitors of metalloproteinases (TIMP) (7, 12, 13). As recently described, DCs express several MMPs including MMP-9, and at least three forms of TIMP (TIMP-1, TIMP-2, and TIMP-3) (14, 15, 16). Moreover, production of active MMP-9 has been shown to contribute to DC migration through ECM (16, 17, 18). Changes in the balance of TIMP and MMP, occurring during maturation of DCs or pathologic conditions, have been implicated in the modulation of DC migratory capacity (15, 16, 17, 18). Growth factors, hormones, inflammatory mediators and cell-matrix interactions regulate the expression of MMP and TIMP (19, 20, 21, 22, 23). In particular, it has been shown that MMP-9 and TIMP-1 are regulated via prostaglandin-dependent and -independent mechanisms in monocytes (24).

PGE2, derived from cyclooxygenase (COX)-dependent metabolism of free arachidonic acid, is overproduced in several malignancies (25, 26, 27) and exerts an important immunomodulatory role in DC differentiation and function (28, 29). PGE2 contributes to DC maturation by up-regulation of costimulatory molecules and modulation of Ag-presenting capacity (28, 29, 30, 31, 32, 33, 34, 35). Recent findings have also shown that PGE2 promotes DC migration in response to chemotactic factors by modulating chemokine receptor expression (31, 32).

PGE2 signals through G protein-coupled E prostanoid (EP) receptors designated EP1, EP2, EP3, and EP4. Expression of these receptors varies in different cell types and mediates the diverse effects of PGE2 and its analogues (36, 37). Despite data describing PGE2 as a key modulator of DC function, a specific role of EP receptors in DC migration through ECM has not been characterized. In the present study, we show that PGE2 enhances TIMP-1 secretion, thus reducing DC migration through ECM.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
In vitro generation and culture of monocyte-derived DC

DCs were generated from PBMC as previously described, with minor modifications (32). The use of leukocyte-enriched buffy coat from healthy donors was approved by the University of California, Los Angeles, Institutional Review Board, and informed consent was obtained from all donors. Briefly, PBMC were obtained from leukocyte-enriched buffy coat from healthy donors (University of California, Los Angeles, Blood Bank) by centrifugation with Ficoll-Paque Plus (Amersham Biosciences, Piscataway, NJ) and the light density fraction from the 42.5–50% interface was recovered. The cells were resuspended in RPMI 1640 (Cellgro; Mediatech, Herndon, VA) supplemented with 20 mM HEPES buffer (Cellgro), 100 U/ml penicillin-streptomycin, 2 mM glutamine (Invitrogen Life Technologies, Grand Island, NY), 2% human serum AB (Gemini Bio-Products, Wooland, CA), and allowed to adhere to tissue culture flasks. After 2 h at 37°C, nonadherent cells were removed and adherent cells were washed twice in PBS (Cellgro). Alternatively, DCs were generated after positive selection of CD14+ cells using microbeads (Miltenyi Biotec, Auburn, CA). Adherent monocytes or CD14+ monocytes were then cultured for 8–10 days in complete RPMI 1640 medium supplemented with 10% human serum AB, 65 ng/ml recombinant human GM-CSF (PeproTech, Rocky Hill, NJ), specific activity, ≥1 x 107 U/mg, and 65 ng/ml recombinant human IL-4 (PeproTech), specific activity, ≥5 x 106 U/mg. To induce maturation, LPS (1 µg/ml; Sigma-Aldrich, St. Louis, MO), TNF-{alpha} (50 ng/ml; PeproTech) or CD40L (1 µg/ml; gift from Dr. G. Zeng, University of California, Los Angeles, CA) were added to DC on day 5 of culture for 48 h. PGE2-treated DC were generated by addition of 16,16-dimethyl-PGE2 (dmPGE2; Cayman Chemicals, Ann Arbor, MI) for 72 h on day 5 of culture for immature DC or on day 7 for LPS and TNF-{alpha}-matured DC. dmPGE2 was used in these studies because of its greater stability in vitro. Preliminary experiments demonstrated that both PGE2 and dmPGE2 had similar effects on TIMP-1 expression (data not shown).

To analyze the effect of tumor-derived-PGE2, on day 5 of culture, DC were incubated for 48 h in supernatants from non-small cell lung cancer cell (NSCLC) lines genetically modified to overexpress COX-2 and therefore secrete high levels of PGE2. At the end-point of the culture, cells exhibited DC morphology and phenotype as assessed by light microscopy and flow cytometry respectively, as previously described (32, 35).

Immunophenotypic analysis of DC by flow cytometry

The DC phenotype was analyzed on day 8 or day 10 of culture. Briefly, all cultured cells were stained with the following mAbs directly labeled with FITC or PE: CD3, CD14, CD40, CD80, CD86, CCR7, HLA-DR (all from BD Biosciences, San Jose, CA), and CD83 (Coulter Immunology, Hialeah, FL). EP2 receptor expression was analyzed in DC treated with or without PGE2 (5 µg/ml) using EP2 receptor rabbit polyclonal Ab (Cayman Chemicals). EP4 receptor expression was analyzed using either an EP4 receptor rabbit polyclonal Ab or a goat anti-human EP4 receptor Ab (Santa Cruz Biotechnology, Santa Cruz, CA). The cells were also stained with the corresponding FITC- or PE-conjugated isotype-matched control Abs (BD Biosciences). In accord with previous reports in the literature, we used CD40L-matured DC and A549 cells as positive controls for CCR7 and EP4, respectively (31, 32, 38). Analysis of fluorescence staining was performed with a Life Science Research flow cytometer (BD Biosciences) and CellQuest software.

TIMP-1 expression analysis by ELISA

Total TIMP-1 secretion was determined from DC culture supernatants using the Biotrak human TIMP-1 ELISA system (Amersham Biosciences) according to the manufacturer’s instructions. Briefly, TIMP-1 standard protein or DC culture supernatants (derived from 0.5 x 105 cells/1.5 ml) were added to each well of an ELISA plate precoated with an anti-TIMP-1 Ab. After 2-h incubation at room temperature, the plate was washed four times and an HRP-conjugated anti-TIMP-1 Ab was added to each well. After 2-h incubation at room temperature, the plate was washed four times to remove all the unbound reagents and the 3,3',5,5'-tetramethylbenzidine substrate buffer was added to each well. Reactions were stopped by adding 1 M sulfuric acid and the OD was read at 450 nm in a Benchmark microplate reader (Bio-Rad, Hercules, CA).

Analysis of total gelatinase B by zymography

Gelatinase B secretion was analyzed by substrate zymography (39), which allows detection of active gelatinase B, the proenzyme form as well as MMP-9 and TIMP-1 complexes (14). Supernatants from DCs (derived from 0.5 x 105 cells/1.5 ml) cultured with or without PGE2 for 72 h were supplemented with an equal volume of 4% SDS-containing zymograph sample buffer (Bio-Rad). The samples were loaded on 10% polyacrylamide gel with gelatin (Bio-Rad). Following electrophoresis the gel was incubated in 2.5% Triton X-100 for 1 h at room temperature and subsequently in a buffer containing 50 mM Tris-HCl (pH 7.5), 200 mM NaCl, 5 mM CaCl2, and 0.02% Brij-35 for 18 h at 37°C. The gel was then stained with Coomassie brilliant blue. After destaining, the proteolytic activity was visualized as clear bands against a blue background. Purified MMP-2 and MMP-9 (Chemicon International, Temecula, CA) were used as control size markers. Densitometry analysis was used to quantify the gelatinolytic activity.

DC migration assay

DC migration through ECM was performed using 24-well Transwell insertsfitted with a 5-µm pore size polycarbonate membrane (Corning, Corning, NY) as previously described (16). Briefly, DCs cultured with or without PGE2 (5 µg/ml) for 72 h, or for 48 h in A549 or H157 cell line culture medium, were harvested and their culture supernatants were harvested to be used in the migration assay. In blocking experiments, DC were preincubated with neutralizing rabbit polyclonal anti-TIMP-1 (10 µg/ml; gift from Dr. L. J. Windsor, Indiana University, Indianapolis, IN) or rabbit IgG (10 µg/ml; Jackson ImmunoResearch Laboratories, West Grove, PA) before addition of dmPGE2 (5 µg/ml) daily for 72 h. A total of 2 x 106 DCs were then labeled with 200 µCi of 51Cr and washed four times in RPMI 1640. A total of 5 x 105 labeled DCs were resuspended in 100 µl of their own culture medium and added to the upper compartment of each insert coated with Growth Factor Reduced Matrigel Matrix (BD Biosciences) diluted in PBS (60 µg/insert).

In some experiments, 0.5 µg/ml recombinant human TIMP-1 (Oncogene Research Products, San Diego, CA) was added to the DC suspension. The recombinant TIMP-1 concentration was chosen based on preliminary experiments, which evaluated TIMP-1 secretion by DC. The lower chambers of the Transwells were filled with 500 µl of RPMI 1640 supplemented with 30% AB serum. Chambers were incubated at 37°C in 5% CO2 for 12–14 h. Migrated cells were recovered from the lower compartment and lysed with PBS containing 0.1% Triton X-100. The radioactivity of the samples was measured with a gamma counter (Beckman Gamma Counter 5500) and expressed as cpm. The DC migratory capacity was determined by dividing the cpm of migrated cells through the ECM filter by the cpm of total input as previously described (16).

To measure DC chemotaxis through Matrigel, 2 x 105 unlabeled immature DCs treated with or without PGE2 and CD40L-matured DCs were resuspended in their culture supernatant and allowed to migrate through the Matrigel-coated inserts against a chemotactic gradient. Secondary lymphoid chemokine (SLC)/CC chemokine ligand (CCL-21/SLC, 250 ng/ml; PeproTech) was used as CCR7 ligand chemoattractant and added to the lower chambers of the Transwells. DC migration was determined after 14-h incubation by flow cytometry as previously described (32).

RT-PCR analysis of EP receptor gene expression

Total RNA was purified from DC cultured with or without PGE2 (5 µg/ml) for 72 h using RNEasy Mini kit (Qiagen, Valencia, CA) according to the manufacturer’s instructions. For the RT-PCR analysis, first-strand cDNA was synthesized from 1.5 µg of total RNA using 200 U of SuperScript II RNase H Reverse Transcriptase (Invitrogen Life Technologies) following the instructions. A one-sixth portion of the cDNA was amplified by PCR using TaqDNA Polymerase (Invitrogen Life Technologies) to examine EP receptor expression (35 cycles of denaturation at 94°C for 30 s, annealing at 55°C for EP1, 62°C for EP2 and EP3 receptors, and at 64°C for EP4 for 30 s and extension at 72°C for 20 or 30 s). The primers used for PCR are listed in Table I (40). The {beta}-actin gene expression was examined as an internal control for the quality of RNA templates.


View this table:
[in this window]
[in a new window]
  		Table I. Nucleotide sequences of the primers used for RT-PCR

 
Stably transduced COX-2 cell lines

NSCLC cells, A549 (human lung adenocarcinoma) and H157 (squamous cell carcinoma), were obtained from American Type Culture Collection (Manassas, VA) and National Cancer Institute, respectively. A 2.0-kb cDNA fragment of human COX-2 (generously provided by Dr. H. Herschman, University of California, Los Angeles, CA) was cloned in the retroviral vector pLNCX (Clontech Laboratories, Palo Alto, CA) in sense (COX-2-S) and antisense (COX-2-AS) orientation as previously described (38, 41). Tumor cells were transduced with high titer-producing COX-2-S and COX-2-AS virus, selected and characterized as previously described. For each cell line, an ~3- to 5-fold higher level of COX-2 expression and PGE2 production was noted in COX-2-S compared with parental. In contrast, COX-2-AS produced less COX-2 and PGE2 than parental. For DC migration assays, 3 x 106 cells of each A549 and H157 cell lines were cultured in RPMI 1640 supplemented with 10% FBS (Gemini Bio-Products), 100 U/ml penicillin/streptomycin, and 2 mM glutamine. After overnight incubation at 37°C in an atmosphere of 5% CO2, culture medium was removed and cells were incubated in RPMI 1640 supplemented with 2% human serum AB. After 72-h incubation, the supernatants were collected and the cells were trypsinized and counted. The supernatants were diluted according to the number of cells. DCs were incubated for 48 h with tumor supernatants collected from each A549 and H157 cultures and the concentration of AB serum was adjusted to 10%.

Statistical analysis

The unpaired two-tailed Student’s t test was used to compare differences in DC TIMP-1 secretion and in DC migration through the extracellular matrix. A p ≤ 0.05 was considered significant. The errors bars shown in all the figures represent the SD of the values.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Generation of immature and mature DC

Studies of DC function have been facilitated by the in vitro generation of DC from monocytes or bone marrow precursors (2). In this study, purified CD14+ monocytes or plastic adherent PBMC were cultured in the presence of GM-CSF and IL-4 for 8–10 days to generate monocyte-derived DC. The resulting cell population resembled immature DC in morphology and phenotype (31, 32, 35): lack of lineage-specific markers (CD14 and CD3, data not shown), high level expression of CD86, HLA-DR, and CD80, and lack of CD83 (Fig. 1). DC maturation was induced on day 5 by addition of either LPS or TNF-{alpha}. Cells showed marked up-regulation of all costimulatory molecules and CD83 (Fig. 1). Immature and mature DCs were exposed to exogenous dmPGE2 for 72 h on day 5 or day 7 of culture, respectively. Following dmPGE2 exposure, the phenotypic features of the immature DC remained unchanged with the exception of a slight up-regulation of CD83 observed in some cases as previously described (32). Up-regulation of CD80 and CD83 and to a lesser extent of CD86 and HLA-DR were noted only in TNF-{alpha} matured DC, following dmPGE2 treatment (Fig. 1).



View larger version (44K):
[in this window]
[in a new window]
  		FIGURE 1. Phenotypic analysis of immature and mature DC cultured in the presence or the absence of PGE2. DCs were matured using LPS (1 µg/ml), TNF-{alpha} (50 ng/ml), or CD40L (1 µg/ml) for 48 h. Immature and LPS- or TNF-{alpha}-matured DCs were cultured with or without dmPGE2 (5 µg/ml) for 72 h. DC phenotype was analyzed by flow cytometry. In the histograms, isotype control (dotted lines) and the surface marker (bold lines) staining are represented. The numbers (upper right) indicate percentage of positive cells. Results are representative of one experiment of three.

 
Lack of expression of CCR7 in PGE2-treated DC

Because PGE2 has been shown to increase CCR7 expression and chemoattraction of monocyte-derived DC (31, 32), we analyzed CCR7 surface expression by flow cytometry in DC after exposure to dmPGE2. Interestingly, under our experimental conditions, CCR7 expression was undetectable in immature DC after dmPGE2 treatment (Fig. 1). When DCs were matured with LPS or TNF-{alpha}, CCR7 was detected at low levels and addition of dmPGE2 at the same time (data not shown) or after DC maturation (Fig. 1) had no strong impact on CCR7 surface expression. In contrast, DC matured with CD40L showed significant up-regulation of CCR7 expression (Fig. 1). In agreement with these results, DC chemotaxis through Matrigel showed that CCL-21/SLC, a CCR7 ligand, increased migration of CD40L-matured DC but not of PGE2-treated immature DC (data not shown). Under our experimental conditions CD40L maturation was required for significant up-regulation of DC CCR7.

PGE2 enhances TIMP-1 secretion in a dose-dependent manner

To determine the effect of PGE2 on TIMP-1 protein expression in DC, immature DCs were cultured with or without dmPGE2 (0.5–5 µg/ml) and TIMP-1 was measured in the supernatant by a specific ELISA detecting both free and complexed TIMP-1. As shown in Fig. 2, PGE2-treated DCs showed a significant increase in TIMP-1 levels compared with control DCs. TIMP-1 secretion was enhanced in response to all of the dmPGE2 concentrations tested with a maximum increase observed between 2 and 5 µg/ml (Fig. 2A). Moreover, the maximum increase in TIMP-1 was detected after 72 h of dmPGE2 exposure (Fig. 2B).



View larger version (17K):
[in this window]
[in a new window]
  		FIGURE 2. PGE2 increases secreted TIMP-1 in immature DC: dose response (A) and time-course (B). Immature DCs were generated from monocytes purified from PBMC and cultured in the presence ({blacksquare}) or in the absence ({square}) of dmPGE2 (5 µg/ml) at different concentrations. TIMP-1 secretion was analyzed from DC culture supernatants by ELISA at different time points. Results are expressed as ratios of values obtained with PGE2-treated DC vs untreated DC and are representative of the average of three experiments. *, Statistically significant difference compared with control DC. Errors bars indicate SD.

 
Modulation of DC TIMP-1 and MMP-9 secretion by PGE2

To assess whether PGE2-induced TIMP-1 has significant biological impact, the balance between TIMP-1 and MMP-9 was analyzed in PGE2-treated and untreated DC. Immature or mature DCs were cultured with or without exogenous dmPGE2 for 72 h and culture supernatants were analyzed for both TIMP-1 and MMP-9 secretion. Total secreted TIMP-1 protein was measured by specific ELISA, and total secreted MMP-9 protein was determined by zymography. The latter method allows the detection of the total amount of enzymatic activity corresponding to the total secretion of gelatinase B (14). Supernatant from both immature and mature DC revealed the presence of gelatinase activity by a single band with an approximate molecular mass of 90,000 Da corresponding to the active enzyme, as previously described (Fig. 3A) (16). Densitometric analysis showed that dmPGE2 did not affect total secreted MMP-9 in immature and LPS-matured DCs compared with their respective controls. Total secreted MMP-9 was increased by dmPGE2 treatment only in TNF-{alpha} matured DCs (Fig. 3A). In contrast, dmPGE2 exposure significantly increased total secreted TIMP-1 in both immature and TNF-{alpha} matured DC, but not in LPS-matured DC compared with their respective controls (Fig. 3B). Thus in response to dmPGE2 TIMP-1 and MMP-9 balance was altered only in immature DC exposed to dmPGE2.



View larger version (30K):
[in this window]
[in a new window]
  		FIGURE 3. Modulation of TIMP-1 and MMP-9 secretion by PGE2 in DC. Immature and mature DCs were cultured with (+) or ({blacksquare}) or without (–) or ({square}) dmPGE2 (5 µg/ml) for 72 h and both TIMP-1 and MMP-9 were analyzed from DC supernatants. A, MMP-9 secretion was analyzed by zymography (top) and gel images were captured by scanning and densitometric analysis was used to quantify the gelatinolytic activity from each sample (bottom). B, Total secreted TIMP-1 was measured by ELISA. Results are expressed as ratio of the control value and represent the average of at least three independent experiments. Errors bars indicate SD. *, Statistically significant differences compared with control.

 
PGE2-mediated enhancement of TIMP-1 production limits the migratory capacity of DC through ECM

MMPs have been identified as key enzymes for the degradation and remodeling of all components of the ECM. As shown in several studies, MMP-9 is implicated in migration of DC through the ECM, and changes in the balance of TIMP and MMP can modulate DC migratory capacity (15, 16, 17, 18, 42). Based on our findings, we speculated that PGE2-dependent enhancement of TIMP-1 production would affect immature DC migratory capacity through tissues and endothelial barriers. Therefore, the migratory capacity of immature DC was analyzed by a Transwell migration assay using ECM-coated filters (16). Because DCs secrete TIMP-1, DC treated with or without dmPGE2 were resuspended in their culture supernatant throughout the migration assay. We found that both PGE2-treated DC and control DC migrated through ECM-coated filters. The average migration for untreated immature DC was ~13%. However, migration of PGE2-treated DC was always reduced, showing an average decline up to 40% compared with control DC (Fig. 4). To determine whether the reduced migration was caused by increased TIMP-1 production, untreated DCs were exposed to recombinant human TIMP-1 (0.5 µg/ml) during the Matrigel assay. Alternatively, DCs were incubated with excess of neutralizing anti-TIMP-1 polyclonal Ab (10 µg/ml) (43) or with the corresponding control IgG (10 µg/ml) before exposure to dmPGE2. The presence of exogenous TIMP-1 reproduced the effect observed with PGE2-treated DC (Fig. 4A), causing a 55% decrease in DC migration through the Matrigel, whereas neutralizing anti-TIMP-1 Ab reversed the PGE2-mediated inhibitory effect on DC migration (Fig. 4B).



View larger version (29K):
[in this window]
[in a new window]
  		FIGURE 4. PGE2-dependent TIMP-1 induction limits DC migratory capacity through ECM. Immature DCs were cultured with or without dmPGE2 (5 µg/ml) for 72 h and migration was assessed using a Matrigel migration assay. In some conditions anti-TIMP-1 neutralizing Ab (10 µg/ml) or rabbit IgG (10 µg/ml) were added 30 min before addition of dmPGE2. 51Cr labeled DCs were resuspended in their own culture medium with or without recombinant TIMP-1 (0.5 µg/ml) and 5 x 105 cells were added in the upper chamber of a Transwell insert-coated with Matrigel. After 12- to 14-h incubation, migration was determined by measuring the radioactivity of migrated cells through the ECM-insert divided by the radioactivity of total input. Results are expressed as the percentage of migration of treated DC to control DC and are representative of the average of at least three independent experiments. Errors bars indicate SD. *, Statistically significant difference compared with control.

 
Tumor derived-PGE2 limits the migratory capacity of DC

PGE2 production is enhanced through up-regulation of COX-2 in a variety of malignancies including lung, breast, and colon cancers (25, 26, 27). Based on the results described earlier, we postulated that tumor-derived PGE2 would induce TIMP-1 secretion in DC and limit DC migration through ECM. Thus, immature DC were cultured for 48 h in the presence of NSCLC cell supernatants obtained from A549 or H157 cells transduced with COX-2-S or COX-2-AS expression vectors, and then used in Matrigel assays.

DC cultured in the presence of A549 or H157 COX-2-S supernatants, which contained high levels of PGE2 (38, 41), showed increased TIMP-1 secretion compared with DC controls (Fig. 5). Consistent with the effect observed when DCs were exposed to exogenous PGE2, DC treated with COX-2-S supernatant showed a greater than 50% decline in migration. DC cultured in A549 or H157 COX-2-S supernatants exhibited significantly lower migration than DC cultured in A549 or H157 COX-2-AS supernatants (Fig. 5). However, DC-treated with COX-2-AS supernatant showed a similar (H157) or reduced (A549) migratory capacity compared with untreated DC, suggesting that a PGE2-independent inhibitory effect of A549 supernatant may also be operative.



View larger version (24K):
[in this window]
[in a new window]
  		FIGURE 5. Tumor-derived PGE2 reduces DC migration through ECM. Immature DCs were cultured with supernatant obtained from the NSCLC A549 or H157 transduced with COX-2-S or COX-2-AS expression vectors for 48 h. DC migration was assessed using a Matrigel migration assay as previously described and TIMP-1 secretion was determined by ELISA. Results are expressed as percentage of migrated cells for DC migration (left panels) and nanograms per milliliter for TIMP-1 secretion (right panels). Results correspond to one representative experiment of three.

 
PGE2-enhanced TIMP-1 secretion is mediated via EP2 receptor signaling

Cellular responses to PGE2 are mediated through the four distinct receptors designated EP1, EP2, EP3, and EP4 (37). To identify the EP receptors responsible for mediating PGE2 signaling in DC, EP receptor expression was analyzed by RT-PCR in PGE2-treated immature and mature DC using {beta}-actin as internal control. Whereas EP1 and EP3 mRNA expression were not detected in DC under our experimental conditions, both EP2 and EP4 receptor transcripts were expressed in DCs when cultured with or without dmPGE2 (Fig. 6A). Analysis of EP2 and EP4 surface expression by flow cytometry, showed that the EP2 receptor was predominantly expressed in DC and it was significantly up-regulated in response to PGE2 only in immature DC (Fig. 6B). To elucidate the functional role of EP receptor subtypes in PGE2-mediated TIMP-1 enhancement, immature DC were treated with EP receptor agonists and antagonists (32, 44). As shown in Fig. 6C, neither sulprostone, an EP3>>EP1 agonist, nor pertussis toxin, which inhibits the effect of Gi coupled receptors such as EP3, caused changes in TIMP-1 secretion, excluding EP1 and EP3 involvement in the PGE2-mediated increase in TIMP-1. These data are consistent with our RT-PCR findings indicating the absence of EP1 and EP3 transcripts in DC. In contrast, forskolin, a pharmacological activator of adenylate cyclase, and cholera toxin, which activates the G{alpha}s subunit of G proteins mimicking Gs coupled receptor signaling (EP2 and EP4), significantly increased TIMP-1 secretion in DC. Given that EP2 and EP4 stimulate cAMP production via Gs proteins, these findings suggest that dmPGE2 signaled TIMP-1 induction through EP2 and/or EP4 receptors. As shown in Fig. 6C, EP2/EP4 receptor agonist PGE1 alcohol and the selective EP2 receptor agonist butaprost also induced a significant increase of TIMP-1 secretion, further supporting the involvement of EP2/EP4 receptors in mediating the dmPGE2 effect in DC. However, DC surface expression of EP receptors suggests a predominant role of EP2 in mediating dmPGE2 signaling.



View larger version (61K):
[in this window]
[in a new window]
  		FIGURE 6. EP receptor expression and signaling in DC and PGE2-treated DC. A, Expression of EP2 and EP4 receptor mRNA in PGE2-treated DC. To analyze EP receptor expression, immature and mature DCs were cultured with or without dmPGE2 (5 µg/ml) for 72 h and RT-PCR was performed on total RNA. Total RNA from NSCLC cells or kidney tissue was used as positive control (+). Negative control (–) corresponds to RT-PCR mixture without RNA samples. Immature DC (1), PGE2-treated immature DC (2), LPS-matured DC (3), TNF-{alpha}-matured DC (4), PGE2-treated LPS-matured DC (5), and PGE2-treated TNF-{alpha}-matured DC (6). B, Surface expression of EP2 and EP4. Immature and LPS or TNF-{alpha}-matured DC cultured with or without dmPGE2 (5 µg/ml) for 72 h were analyzed by flow cytometry for surface expression of EP2 and EP4 receptors. B, Histograms show surface marker staining (bold lines) and isotype control (dotted lines). The numbers (upper right) indicate percentage of positive cells. Results are representative of one experiment of three. C, Effect of EP receptor agonists and antagonists on DC TIMP-1 secretion. DCs were cultured with dmPGE2 (5 µg/ml), PGE1 alcohol (15 µM), forskolin (20 µM), cholera toxin (1 µg/ml), sulprostone (1 µM), butaprost (1 µM), pertussis toxin (50 ng/ml), or pertussis toxin (50 ng/ml) + dmPGE2 (5 µg/ml) for 72 h. TIMP-1 was analyzed by ELISA in the supernatant of the cells and the results were expressed as relative TIMP-1 increase. Results are representative of the average of three independent experiments. Error bars indicate SD. *, Statistically significant difference compared with control DC.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
DCs are the most efficient type of migrating APC, playing an essential role in the initiation of immune responses (1, 2). To fulfill their effector function, DCs migrate through endothelial barriers and tissues. Although recent studies have shown that some of the mechanisms underlying DC migration are controlled by MMP and TIMP (16, 17, 18), little is known about TIMP and MMP regulation of DC migratory capacity.

PGE2, a major product of arachidonic acid metabolism, is engaged in a complex regulatory network that modulates the actions of immune cells during inflammation and tumorigenesis (36, 45). In vitro studies have shown that PGE2 affects maturation and differentiation of cultured DC (28, 29, 30, 32, 33, 34, 35). Moreover, PGE2 has been implicated in the regulation of MMP and TIMP in various cell types, including monocytes (24, 46, 47). Thus, we hypothesized that PGE2 would affect DC migratory capacity by modulating MMP and TIMP secretion.

The current study demonstrates that exposure of immature human monocyte-derived DC to exogenous PGE2 enhanced TIMP-1 secretion but not MMP-9 production, altering the balance between TIMP-1 and MMP-9. In contrast, addition of PGE2 to TNF-{alpha}-stimulated DC induced secretion of both MMP-9 and TIMP-1. Addition of PGE2 to LPS-matured DC did not significantly induce TIMP-1 and MMP-9 secretion. Thus, the overall balance in TIMP-1 and MMP-9 remained unchanged in response to PGE2 in DC matured by either stimulus.

Cellular responses to PGE2 are mediated via four different prostanoid receptors. In contrast to murine DC, which express all four EP receptors (48), and in agreement with previous reports in human monocyte-derived DC (32), we found that DC expressed only EP2 and EP4 as measured at the RNA level. Because flow cytometry analysis revealed that both immature and mature DC express similar levels of EP2 and EP4 surface receptors, we cannot correlate PGE2-mediated enhancement of TIMP-1 production in immature DC to a differential expression of EP2 and/or EP4.

In addition, we found that treatment of immature DC with EP receptor agonists and antagonists showed that only EP2/EP4 agonists replicated the effects observed with exogenous PGE2. However, because immature DC predominantly expressed EP2, as revealed by flow cytometry, PGE2-enhanced TIMP-1 secretion is likely to occur via EP2 receptor signaling. Recent reports have described the importance of EP4 receptor in mediating PGE2 cell signaling: in murine Langerhans cells PGE2 appeared to promote migration to regional lymph nodes and in an NK cell line to regulate MMP expression through EP4-dependent mechanisms (49, 50). In the same line of evidence, we also previously described a PGE2 mediated-EP4 receptor-dependent induction of MMP-2 and CD44 in NSCLC cells (41). Together, our and other studies (51, 52) suggest a distinct role of PGE2 operating via different signaling mechanisms in a cell type-dependent and species-specific manner.

Our results also show that exposure of immature DC to PGE2 (5 µg/ml) significantly up-regulated DC surface expression of EP2 receptors. Recently, Scandella et al. (32) found that a lower concentration of PGE2 (1 µg/ml) down-regulated EP2 receptor mRNA level (32). In agreement with this study, when we cultured immature DC with the same lower PGE2 concentration we obtained similar results, suggesting a dose-dependent modulation of EP receptors by PGE2 in immature DC.

As sentinels of the immune system, immature DC capture and process Ags at tissue sites (1). Simultaneously DCs undergo a process of maturation by up-regulation of costimulatory molecules while migrating to lymphoid tissues. Several studies have shown that PGE2 promotes invasion of a variety of cancer cells, such as lung, breast, and colorectal carcinoma cells (38, 40, 41, 53, 54). Interestingly, we found that PGE2 decreased the migratory capacity of immature DC through the ECM, thus depending on the cell type, PGE2 can act either to promote or inhibit migration.

Two recent studies have shown that PGE2-containing stimuli promote DC migration through PGE2-dependent up-regulation of CCR7 (31, 32). In these reports, DC migration was analyzed in response to specific chemoattractants (CCL-19 and CCL-21). Accordingly, when we analyzed DC migration through ECM toward CCL-21/SLC, we did not obtain increased immature DC migration through Matrigel (data not shown), consistent with the lack of surface CCR7 expression detected in immature DC treated with or without PGE2 (Fig. 1). Substantial differences in the culture conditions (culture medium and PGE2 formulation) and the experimental design (Matrigel coated inserts) used to assess DC migration may explain these dissimilar results. Nonetheless, all these findings highlight the contribution of distinct mechanisms in the complex process of DC migration.

Tight regulation of MMP and TIMP expression is crucial in the degradation of ECM, which occurs in several physiological and pathological processes (9, 23). Any imbalance in favor of MMP inhibitors can lead to deficient cleavage of matrix components and to fibrotic processes, whereas increases in enzymatic activity will result in tissue destruction or cell migration. Recent studies have shown that MMP-9 participates in the migration of murine Langerhans cells in vitro and in vivo and that MMP and TIMP expression define the migratory characteristics of DC (15, 16, 17). In this study, we show that neutralizing anti-TIMP-1 polyclonal Ab (43) was able to reverse the inhibitory effect of PGE2 on DC migration, and addition of recombinant TIMP-1 could substitute for exposure to exogenous PGE2. We conclude that the PGE2-dependent increase in TIMP-1 secretion limits DC migration through ECM. Because PGE2 increases TIMP-1 via EP2/EP4 receptors, future studies will address whether migration through ECM is also regulated by EP2/EP4 signaling. Although our study has focused on TIMP-1 and MMP-9 imbalance in PGE2-treated DC, it is possible that TIMP-1-dependent DC migratory inhibition observed may be also related to other MMPs. Indeed TIMP-1 binds to other MMPs secreted by monocyte-derived DC, such as MMP-1 and MMP-2 (15, 16), the latter being implicated in Langerhans cells migration (42).

It is well established that PGE2 production is enhanced through up-regulation of COX-2 in various cancer cells including NSCLC (45). As immune cells present in the tumor environment, DCs are potentially exposed to the effects of tumor-PGE2 overproduction (29). Our findings indicate that NSCLC-derived PGE2, increased TIMP-1 secretion and limited immature DC migration through ECM. Therefore tumor derived-PGE2, by inducing TIMP-1, could affect the migratory capacity of DC at the tumor site, leading to abnormal sequestration of immature DC within neoplastic tissues. This hypothesis is supported by Bell et al. (55) who observed an increase of immature DC within breast carcinoma tissues compared with normal breast epithelium. In contrast, mature DC were exclusively detected in peritumoral areas of breast carcinoma tissues, supporting the hypothesis that PGE2 would not affect migratory capacity of mature DC because TIMP-1 and MMP-9 balance remained unchanged. Additional PGs may be present in the tumor environment and impact DC migration. For example, PGD2 has been detected in human tumors (56) and has also been implicated in inhibition of DC migration (57, 58). Further studies will be required to define the role of PGD2 in monocyte-derived DC migratory capacity through ECM.

The current study has focused on the effect of PGE2-mediated TIMP-1 and MMP-9 imbalance on the migratory capacity of immature DC through ECM. Although the best-known functions of MMP are the degradation and remodeling of all components of the ECM, other substrates are also degraded by MMP. For example, MMP-9 is able to cleave proinflammatory cytokines and chemokines such as IL-1 and IL-8, resulting in either activation or inactivation of these molecules (59). Moreover, recent studies have demonstrated that MMP-9 is able to cleave myelin basic protein and collagenase type II with exposure of immunodominant epitopes to autoreactive T cells, potentially leading to autoimmune reactions (14, 59). Therefore, the PGE2-dependent enhancement of TIMP-1 secretion by DC could have profound consequences for regulation of immune responses. In addition, increased TIMP-1 has recently been shown to contribute to tumorigenesis, by increasing proliferation and genomic instability in keratinocytes (60). Thus, PGE2-dependent enhancement of DC TIMP-1 secretion could promote tumorigenesis by both immune-dependent and -independent pathways.

This is the first report of PGE2-mediated modulation of TIMP-1 impacting DC migration through ECM. The current study provides further evidence that PGE2 may play an important immunosuppressive role in cancer by compromising DC trafficking. Tumor-derived PGE2 may modify DC trafficking by enhancing TIMP-1 production and thus reducing ECM migration.


   Acknowledgments
 
Flow cytometry was performed in the University of California, Los Angeles, Jonsson Comprehensive Cancer Center and Center for AIDS Research Flow Cytometry Core Facility that is supported by National Institutes of Health Award Grants CA-16042 and AI-28697, by the Jonsson Cancer Center and AIDS Institute, and by the David Geffen School of Medicine, University of California, Los Angeles, CA. We are grateful to Dr. L. Jack Windsor, Indiana University (Indianapolis, IN) for providing neutralizing anti-TIMP-1 polyclonal Ab (Ab 2315) and to Dr. Gang Zeng (University of California, Los Angeles, CA) for providing CD40L trimer.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by the University of California, Los Angeles, Specialized Programs of Research Excellence in Lung Cancer, National Institutes of Health Grants P50 CA90388 and RO1 CA85686, the American Lung Association, Merit Review Research Funds from the Department of Veterans Affairs, and the Tobacco-Related Disease Research Program of the University of California, Los Angeles, CA. Back

2 F.E.B. and N.H.-V. contributed equally to this study. Back

3 Address correspondence and reprint requests to Dr. Steven M. Dubinett, Lung Cancer Research Program, Division of Pulmonary and Critical Care Medicine, David Geffen School of Medicine, University of California, 37-131 Center for Health Sciences, 10833 Le Conte Avenue, Los Angeles, CA 90095. E-mail address: sdubinett{at}mednet.ucla.edu Back

4 Abbreviations used in this paper: DC, dendritic cell; dmPGE2, 16,16-dimethyl PGE2; EP, E prostanoid receptor; NSCLC, non-small cell lung cancer; MMP, metalloproteinase; TIMP, tissue inhibitors of metalloproteinase; ECM, extracellular matrix; SLC, secondary lymphoid chemokine; COX, cyclooxygenase. Back

Received for publication November 17, 2003. Accepted for publication August 19, 2004.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Banchereau, J., R. M. Steinman. 1998. Dendritic cells and the control of immunity. Nature 392:245.[Medline]
  2. Banchereau, J., F. Briere, C. Caux, J. Davoust, S. Lebecque, Y. J. Liu, B. Pulendran, K. Palucka. 2000. Immunobiology of dendritic cells. Annu. Rev. Immunol. 18:767.[Medline]
  3. Cyster, J. G.. 1999. Chemokines and cell migration in secondary lymphoid organs. Science 286:2098.[Abstract/Free Full Text]
  4. Dieu-Nosjean, M. C., A. Vicari, S. Lebecque, C. Caux. 1999. Regulation of dendritic cell trafficking: a process that involves the participation of selective chemokines. J. Leukocyte Biol. 66:252.[Abstract]
  5. Caux, C., S. Ait-Yahia, K. Chemin, O. de Bouteiller, M. C. Dieu-Nosjean, B. Homey, C. Massacrier, B. Vanbervliet, A. Zlotnik, A. Vicari. 2000. Dendritic cell biology and regulation of dendritic cell trafficking by chemokines. Springer Semin. Immunopathol. 22:345.[Medline]
  6. Sozzani, S., P. Allavena, A. Vecchi, A. Mantovani. 1999. The role of chemokines in the regulation of dendritic cell trafficking. J. Leukocyte Biol. 66:1.[Abstract]
  7. Brown, P. D., R. Giavazzi. 1995. Matrix metalloproteinase inhibition: a review of anti-tumour activity. Ann. Oncol. 6:967.[Abstract/Free Full Text]
  8. Faveeuw, C., G. Preece, A. Ager. 2001. Transendothelial migration of lymphocytes across high endothelial venules into lymph nodes is affected by metalloproteinases. Blood 98:688.[Abstract/Free Full Text]
  9. Vu, T. H., Z. Werb. 2000. Matrix metalloproteinases: effectors of development and normal physiology. Genes Dev. 14:2123.[Free Full Text]
  10. Emonard, H., J. A. Grimaud. 1990. Matrix metalloproteinases: a review. Cell. Mol. Biol. 36:131.[Medline]
  11. Timpl, R.. 1989. Structure and biological activity of basement membrane proteins. Eur. J. Biochem. 180:487.[Medline]
  12. Brew, K., D. Dinakarpandian, H. Nagase. 2000. Tissue inhibitors of metalloproteinases: evolution, structure and function. Biochim. Biophys. Acta 1477:267.[Medline]
  13. Nagase, H., J. F. Woessner, Jr. 1999. Matrix metalloproteinases. J. Biol. Chem. 274:21491.[Free Full Text]
  14. Bartholome, E. J., I. Van Aelst, E. Koyen, R. Kiss, F. Willems, M. Goldman, G. Opdenakker. 2001. Human monocyte-derived dendritic cells produce bioactive gelatinase B: inhibition by IFN-{beta}. J. Interferon Cytokine Res. 21:495.[Medline]
  15. Kouwenhoven, M., V. Ozenci, A. Tjernlund, M. Pashenkov, M. Homman, R. Press, H. Link. 2002. Monocyte-derived dendritic cells express and secrete matrix-degrading metalloproteinases and their inhibitors and are imbalanced in multiple sclerosis. J. Neuroimmunol. 126:161.[Medline]
  16. Osman, M., M. Tortorella, M. Londei, S. Quaratino. 2002. Expression of matrix metalloproteinases and tissue inhibitors of metalloproteinases define the migratory characteristics of human monocyte-derived dendritic cells. Immunology 105:73.[Medline]
  17. Kobayashi, Y., M. Matsumoto, M. Kotani, T. Makino. 1999. Possible involvement of matrix metalloproteinase-9 in Langerhans cell migration and maturation. J. Immunol. 163:5989.[Abstract/Free Full Text]
  18. Hollender, P., D. Ittelet, F. Villard, J. C. Eymard, P. Jeannesson, J. Bernarrd. 2002. Active matrix metalloproteinase-9 in and migration pattern of dendritic cells matured in clinical grade culture conditions. Immunobiology 206:441.[Medline]
  19. Borden, P., R. A. Heller. 1997. Transcriptional control of matrix metalloproteinases and the tissue inhibitors of matrix metalloproteinases. Crit. Rev. Eukaryotic Gene Expression 7:159.[Medline]
  20. Janowska-Wieczorek, A., L. A. Marquez, J. M. Nabholtz, M. L. Cabuhat, J. Montano, H. Chang, J. Rozmus, J. A. Russell, D. R. Edwards, A. R. Turner. 1999. Growth factors and cytokines upregulate gelatinase expression in bone marrow CD34+ cells and their transmigration through reconstituted basement membrane. Blood 93:3379.[Abstract/Free Full Text]
  21. Johnatty, R. N., D. D. Taub, S. P. Reeder, S. M. Turcovski-Corrales, D. W. Cottam, T. J. Stephenson, R. C. Rees. 1997. Cytokine and chemokine regulation of proMMP-9 and TIMP-1 production by human peripheral blood lymphocytes. J. Immunol. 158:2327.[Abstract]
  22. Kossakowska, A. E., D. R. Edwards, C. Prusinkiewicz, M. C. Zhang, D. Guo, S. J. Urbanski, T. Grogan, L. A. Marquez, A. Janowska-Wieczorek. 1999. Interleukin-6 regulation of matrix metalloproteinase (MMP-2 and MMP-9) and tissue inhibitor of metalloproteinase (TIMP-1) expression in malignant non-Hodgkin’s lymphomas. Blood 94:2080.[Abstract/Free Full Text]
  23. Parks, W. C., S. D. Shapiro. 2001. Matrix metalloproteinases in lung biology. Respir. Res. 2:10.[Medline]
  24. Zhang, Y., K. McCluskey, K. Fujii, L. M. Wahl. 1998. Differential regulation of monocyte matrix metalloproteinase and TIMP-1 production by TNF-{alpha}, granulocyte-macrophage CSF, and IL-1{beta} through prostaglandin-dependent and -independent mechanisms. J. Immunol. 161:3071.[Abstract/Free Full Text]
  25. Huang, M., S. Sharma, J. T. Mao, S. M. Dubinett. 1996. Non-small cell lung cancer-derived soluble mediators and prostaglandin E2 enhance peripheral blood lymphocyte IL-10 transcription and protein production. J. Immunol. 157:5512.[Abstract]
  26. Dannenberg, A. J, R. N. DuBois. 2003. COX-2 a new target for cancer prevention and treatment. In Progress in Experimental Tumor Research vol. 37: Karger, Cambridge, Mass.
  27. Turini, M. E, R. N. DuBois. 2002. Cyclooxygenase-2: a therapeutic target. Annu. Rev. Med. 53:35.[Medline]
  28. Harizi, H., M. Juzan, V. Pitard, J. F. Moreau, N. Gualde. 2002. Cyclooxygenase-2-issued prostaglandin E2 enhances the production of endogenous IL-10, which down-regulates dendritic cell functions. J. Immunol. 168:2255.[Abstract/Free Full Text]
  29. Sharma, S., M. Stolina, S. C. Yang, F. Baratelli, J. F. Lin, K. Atianzar, J. Luo, L. Zhou, Y. Lin, M. Huang, et al 2003. Tumor cyclooxygenase 2-dependent suppression of dendritic cell function. Clin. Cancer Res. 3:961.
  30. Whittaker, D. S., K. S. Bahjat, L. L. Moldawer, M. J. Clare-Salzler. 2000. Autoregulation of human monocyte-derived dendritic cell maturation and IL-12 production by cyclooxygenase-2-mediated prostanoid production. J. Immunol. 165:4298.[Abstract/Free Full Text]
  31. Luft, T., M. Jefford, P. Luetjens, T. Toy, H. Hochrein, K. A. Masterman, C. Maliszewski, K. Shortman, J. Cebon, E. Maraskovsky. 2002. Functionally distinct dendritic cell (DC) populations induced by physiologic stimuli: prostaglandin E2 regulates the migratory capacity of specific DC subsets. Blood 100:1362.[Abstract/Free Full Text]
  32. Scandella, E., Y. Men, S. Gillessen, R. Forster, M. Groettrup. 2002. Prostaglandin E2 is a key factor for CCR7 surface expression and migration of monocyte-derived dendritic cells. Blood 100:1354.[Abstract/Free Full Text]
  33. Kalinski, P., C. M. Hilkens, A. Snijders, F. G. Snijdewint, M. L. Kapsenberg. 1997. IL-12-deficient dendritic cells, generated in the presence of prostaglandin E2, promote type 2 cytokine production in maturing human naive T helper cells. J. Immunol. 159:28.[Abstract]
  34. Wu, C. Y., K. Wang, J. F. McDyer, R. A. Seder. 1998. Prostaglandin E2 and dexamethasone inhibit IL-12 receptor expression and IL-12 responsiveness. J. Immunol. 161:2723.[Abstract/Free Full Text]
  35. Rieser, C., G. Bock, H. Klocker, G. Bartsch, M. Thurnher. 1997. Prostaglandin E2 and tumor necrosis factor {alpha} cooperate to activate human dendritic cells: synergistic activation of interleukin 12 production. J. Exp. Med. 186:1603.[Abstract/Free Full Text]
  36. Tilley, S. L., T. M. Coffman, B. H. Koller. 2001. Mixed messages: modulation of inflammation and immune responses by prostaglandins and thromboxanes. J. Clin. Invest. 108:15.[Medline]
  37. Breyer, R. M., C. K. Bagdassarian, S. A. Myers, M. D. Breyer. 2001. Prostanoid receptors: subtypes and signaling. Annu. Rev. Pharmacol. Toxicol. 41:661.[Medline]
  38. Dohadwala, M., R. J. Batra, J. Luo, Y. Lin, K. Krysan, M. Pold, S. Sharma, S. M. Dubinett. 2002. Autocrine/paracrine prostaglandin E2 produiction by non-small cell lung cancer cells regulates matrix metalloproteinase-2 and CD44 in cyclopxygenase-2-dependent invasion. J. Biol. Chem. 277:50828.[Abstract/Free Full Text]
  39. Masure, S., A. Billiau, J. Van Damme, G. Opdenakker. 1990. Human hepatoma cells produce an 85 kDa gelatinase regulated by phorbol 12-myristate 13-acetate. Biochim. Biophys. Acta 1054:317.[Medline]
  40. Sheng, H., J. Shao, M. K. Washington, R. N. DuBois. 2001. Prostaglandin E2 increases growth and motility of colorectal carcinoma cells. J. Biol. Chem. 276:18075.[Abstract/Free Full Text]
  41. Dohadwala, M., J. Luo, L. Zhou, Y. Lin, G. Dougherty, S. Sharma, M. Huang, M. Pold, R. K. Batra, S. M. Dubinett. 2001. Non-small cell lung cancer cyclooxygenase-2 dependent invasion is mediated by CD44. J. Biol. Chem. 276:20809.[Abstract/Free Full Text]
  42. Ratzinger, G., P. Stoitzner, S. Ebner, M. B. Lutz, G. T. Layton, C. Rainer, R. M. Senior, J. M. Shipley, P. Fritsch, G. Schuler, N. Romani. 2002. Matrix metalloproteinases 9 and 2 are necessary for the migration of Langerhans cells and dermal dendritic cells from human and murine skin. J. Immunol. 168:4361.[Abstract/Free Full Text]
  43. Bodden, M. K., G. J. Harber, B. Birkedal-Hansen, L. J. Windsor, N. C. M. Caterina, J. A. Angler, H. Birkedal-Hansen. 1994. Functional domains of human TIMP-1 (tissue inhibitor of metalloproteinases). J. Biol. Chem. 269:18943.[Abstract/Free Full Text]
  44. Abramovitz, M., M. Adam, Y. Boie, M. Carriere, D. Denis, C. Godbout, S. Lamontagne, C. Rochette, N. Sawyer, N. M. Tremblay, et al 2000. The utilization of recombinant prostanoid receptors to determine the affinities and selectivities of prostaglandins and related analogs. Biochim. Biophys. Acta 1483:285.[Medline]
  45. Huang, M., M. Stolina, S. Sharma, J. T. Mao, L. Zhu, P. W. Miller, J. Wollman, H. Herschman, S. M. Dubinett. 1998. Non-small cell lung cancer cyclooxygenase-2-dependent regulation of cytokine balance in lymphocytes and macrophages: up-regulation of interleukin 10 and down-regulation of interleukin 12 production. Cancer Res. 58:1208.[Abstract/Free Full Text]
  46. Liu, C. M., C. Y. Hong, C. T. Shun, J. S. Wang, T. Y. Hsiao, C. C. Wang, S. K. Lin. 2001. Matrix metalloproteinase-1 and tissue inhibitor of metalloproteinase-1 gene expressions and their differential regulation by proinflammatory cytokines and prostaglandin in nasal polyp fibroblasts. Ann. Otol. Rhinol. Laryngol. 110:1129.[Medline]
  47. Tung, J. T., C. E. Arnold, L. H. Alexander, V. Yuzbasiyan-Gurkan, P. J. Venta, D. W. Richardson, J. P. Caron. 2002. Evaluation of the influence of prostaglandin E2 on recombinant equine interleukin-1{beta}-stimulated matrix metalloproteinases 1, 3, and 13 and tissue inhibitor of matrix metalloproteinase 1 expression in equine chondrocyte cultures. Am. J. Vet. Res. 63:987.[Medline]
  48. Harizi, H., C. Grosset, N. Gualde. 2003. Prostaglandin E2 modulates dendritic cell function via EP2 and EP4 receptor subtypes. J. Leukocyte Biol. 73:756.[Abstract/Free Full Text]
  49. Kabashima, K., D. Sakata, M. Nagamachi, Y. Miyachi, K. Inaba, S. Narumiya. 2003. Prostaglandin E2-EP4 signaling initiates skin immune responses by promoting migration and maturation of Langerhans cells. Nat. Med. 9:744.[Medline]
  50. Zeng, L., S. An, E. J. Goetzl. 1996. Selective regulation of RNK-16 cell matrix metalloproteinases by the EP4 subtype of prostaglandin E2 receptor. Biochemistry 35:7159.[Medline]
  51. Harris, S. R., J. Padilla, L. Koumas, D. Ray, R. P. Phipps. 2002. Prostaglandins as modulator of immunity. Trends Immunol. 23:144.[Medline]
  52. Yang, L., N. Yamagata, R. Yadav, S. Brandon, R. L. Courtney, J. D. Morrow, Y. Shyr, M. Boothby, S. Joyce, D. P. Carbone, R. M. Breyer. 2003. Cancer-associated immunodeficiency and dendritic cell abnormalities mediated by the prostaglandin EP2 receptor. J. Clin. Invest. 111:727.[Medline]
  53. Matsumoto-Taniura, N., K. Matsumoto, T. Nakamura. 1999. Prostaglandin production in mouse mammary tumour cells confers invasive growth potential by inducing hepatocyte growth factor in stromal fibroblasts. Br. J. Cancer 81:194.[Medline]
  54. Rozic, J. G., C. Chakraborty, P. K. Lala. 2001. Cyclooxygenase inhibitors retard murine mammary tumor progression by reducing tumor cell migration, invasiveness and angiogenesis. Int. J. Cancer. 93:497.[Medline]
  55. Bell, D., P. Chomarat, D. Broyles, G. Netto, G. M. Harb, S. Lebecque, J. Valladeau, J. Davoust, K. A. Palucka, J. Banchereau. 1999. In breast carcinoma tissue, immature dendritic cells reside within the tumor, whereas mature dendritic cells are located in peritumoral areas. J. Exp. Med. 190:1417.[Abstract/Free Full Text]
  56. Paoletti, P., C. Chiabrando, P. Gaetani, M. G. Castelli, G. Butti, L. Martelli, M. Rolli. 1989. Prostaglandins in human brain tumors. J. Neurosurg. Sci. 33:65.[Medline]
  57. Hammad, H., H. J. de Heer, T. Soullie, H. C. Hoogsteden, F. Trottein, B.N. Lambrecht. 2003. Prostaglandin D2 inhibits airway dendritic cell migration and function in steady state conditions by selective activation of the D prostanoid receptor I.. J. Immunol. 171:3936.[Abstract/Free Full Text]
  58. Gosset, P., F. Bureau, V. Angeli, M. Pichavant, C. Faveeuw, A. B. Tonnel, F. Trottein. 2003. Prostaglandin D2 affects maturation of human monocyte-derived dendritic cells: consequence on the polarization of naive Th cells. J. Immunol. 170:4943.[Abstract/Free Full Text]
  59. Opdenakker, G., P. E. Van den Steen, J. Van Damme. 2001. Gelatinase B: a tuner and amplifier of immune functions. Trends Immunol 22:571.[Medline]
  60. Rhee, J. S., R. Diaz, L. Korets, J. G. Hodgson, L. M. Coussens. 2004. TIMP-1 alters susceptibility to carcinogenesis. Cancer Res. 64:952.[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 J. Nutr.Home page
D. E. Lackey, S. L. Ashley, A. L. Davis, and K. A. Hoag
Retinoic Acid Decreases Adherence of Murine Myeloid Dendritic Cells and Increases Production of Matrix Metalloproteinase-9
J. Nutr., August 1, 2008; 138(8): 1512 - 1519.
[Abstract] [Full Text] [PDF]

Home page
 J. Leukoc. Biol.Home page
M. Lehner, A. Stilper, P. Morhart, and W. Holter
Plasticity of dendritic cell function in response to prostaglandin E2 (PGE2) and interferon-{gamma} (IFN-{gamma})
J. Leukoc. Biol., April 1, 2008; 83(4): 883 - 893.
[Abstract] [Full Text] [PDF]

Home page
 J. Exp. Med.Home page
G. Rotta, G. Matteoli, E. Mazzini, P. Nuciforo, M. P. Colombo, and M. Rescigno
Contrasting roles of SPARC-related granuloma in bacterial containment and in the induction of anti-Salmonella typhimurium immunity
J. Exp. Med., March 17, 2008; 205(3): 657 - 667.
[Abstract] [Full Text] [PDF]

Home page
 BloodHome page
J.-H. Yen, T. Khayrullina, and D. Ganea
PGE2-induced metalloproteinase-9 is essential for dendritic cell migration
Blood, January 1, 2008; 111(1): 260 - 270.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Respir. Cell Mol. Bio.Home page
S. Razani-Boroujerdi and M. L. Sopori
Early Manifestations of NNK-Induced Lung Cancer: Role of Lung Immunity in Tumor Susceptibility
Am. J. Respir. Cell Mol. Biol., January 1, 2007; 36(1): 13 - 19.
[Abstract] [Full Text] [PDF]

Home page
 J. Leukoc. Biol.Home page
N. L. Webster and S. M. Crowe
Matrix metalloproteinases, their production by monocytes and macrophages and their potential role in HIV-related diseases
J. Leukoc. Biol., November 1, 2006; 80(5): 1052 - 1066.
[Abstract] [Full Text] [PDF]