Published online before print November 7, 2005
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* Experimental Transplantation and Immunology Branch, Center for Cancer Research, National Cancer Institute, and
Laboratory of Neurochemistry, National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, Maryland
1 Correspondence: Experimental Transplantation and Immunology Branch, National Cancer Institute, Building 10, Room 12C205, 10 Center Drive 1907, Bethesda, MD 20892. E-mail: Nakayamt{at}mail.nih.gov
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60%. Agonists of PGE2 receptor subtypes revealed that the EP1 and EP3 receptors can independently mediate MCP-1 release from MCs. These observations identify PGE2-induced MCP-1 release from MCs as a pathway underlying inflammation-associated angiogenesis and extend current understanding of the activities of PGE2.
Key Words: inflammation • chemokine • angiogenesis
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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(TNF-), which serve to mediate acute allergic reactions [4
, 5
]. MCs are also found in the connective tissue of many organs and localize around blood and lymphatic microvessels [6
]. The broad tissue distribution of MCs in proximity to capillary vessels and their prominence in inflammatory infiltrates has suggested a wider range of MC activities than previously thought. Recently, genetic and functional experiments have provided strong evidence for a critical role of MCs in promoting tumor angiogenesis and tumor growth [7
, 8
]. Unlike other cells within the inflammatory infiltrate, MCs are a rich source of factors that regulate angiogenesis, including heparin [9 ], histamine [3 ], vascular endothelial growth factor (VEGF)-A [10 ], basic fibroblast growth factor (bFGF) [11 ], angiopoietin (Ang)-1 [8 ], Ang-2 [12 ], interleukin (IL)-8 [13 ], matrix metalloproteinases [14 ], and monocyte chemoattractant protein (MCP)-1 [15 16 17 ].
MCP-1 (CC chemokine ligand 2), a CC chemokine that serves as a major monocyte chemoattractant [18 19 20 ], can exert direct, proangiogenic functions by locally recruiting endothelial cells that express the MCP-1 receptor CC chemokine receptor 2 (CCR2) [21 ]. It can also promote angiogenesis indirectly, through chemotaxis of monocytes and macrophages, which are themselves proangiogenic [22 23 24 ].
Prostaglandin E2 (PGE2), an arachidonic acid (AA) metabolite generated by the cyclooxygenase enzymes COX-1 and COX-2 in macrophages, fibroblasts, and other cells, is recognized as a potent, proinflammatory mediator, which exerts its biological activities by interaction with one or a combination of subtype receptors designated EP1, EP2, EP3, and EP4, variously induced by proinflammatory signals [25 ]. In some inflammatory conditions, a direct relationship is detected between inflammatory disease activity and levels of PGE2 in tissues. COX-2 inhibitors, which can reduce PGE2 production, are effective at reducing inflammation and inflammatory disease progression [26 ]. In a murine model of pristane-induced plasmacytomagenesis, indomethacine, a COX-2 inhibitor, which diminishes PGE2 production, suppresses cancer development at the site of chronic inflammation [27 28 29 30 ].
MCs often participate in inflammatory processes associated with PGE2 release and are detected in close proximity to PGE2-producing fibroblasts [31 ] and macrophages [32 ]. PGE2 has been reported to favor MC development from murine spleen cell precursors [33 ] and from human umbilical cord blood mononuclear cells [34 ] and to promote or potentiate IL-6 production in murine bone marrow-derived MCs (BMMCs), acting through EP1 and/or EP3 receptors [35 ]. However, little is known about the effects of PGE2 on MC release of proangiogenic factors. Recently, PGE2 was reported to promote VEGF-A production in human cord blood mononuclear cell-derived MCs via activation of the EP2 receptor [36 ]. In this study, we have used well-characterized populations of primary murine MCs derived from the bone marrow (BMMC) and spleen {spleen-derived MC (SPMC) [15 , 37 ]} to evaluate the effect of PGE2 on MC expression and release of proangiogenic factors.
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-PGE2, and sulprostone were from Cayman Chemical (Ann Arbor, MI). Phorbol 12-myristate 13 acetate (PMA), calcium ionophore A23187, and dexamethasone (DM) were obtained from Sigma-Aldrich (St. Louis, MO). Cyclosporine A (CsA) was purchased from Novartis Pharmaceuticals (Basel, Switzerland). RPMI 1640 and fetal bovine serum (FBS) were purchased from Biofluids (Rockville, MD). BMMCs and SPMCs, obtained from C57BL/6 mice [National Cancer Institute (NCI), Frederick, MD], prepared as we described previously [8
], were cultured with RPMI-1640 medium containing 10% FBS, 50 µM 2-mercaptoethanol, and 10 ng/mL IL-3 (Peprotech Corp, Seattle, WA) as described previously [8
, 38
, 39
]. After 4–5 weeks of culture, more than 98% of cells were identified as MC by toluidine blue staining or flow cytometry for surface c-kit and Fc receptor for immunoglobulin E (IgE; Fc
R)I [8
, 40
]. WEHI-3 cells (a gift of Dr. Masahi Narazaki, Osaka University, Japan) were cultured in RPMI-1640 medium containing 10% FBS.
RNA preparation and reverse transcriptase-polymerase chain reaction (RT-PCR) analysis
Total RNA was extracted from BMMCs using TRIzol reagent (Molecular Research Center, Cincinnati, OH). cDNA was synthesized from 5 µg total RNA using the SuperScript preamplification system (Gibco-BRL, Grand Island, NY), followed by DNase I treatment (Invitrogen, Carlsbad, CA). Semiquantitative RT-PCR was carried out to evaluate PGE2-induced modulation of MCP-1 mRNA. The amount of cDNA used for each amplification reaction was based on the results of PCR for glyceraldehyde 3-phosphate dehdyrogenase (GAPDH), showing equivalent amounts of product amplified from all samples. The number of amplification cycles was determined experimentally for each primer pair to fit the linear part of the sigmoid curve, reflecting the relationship between the number of amplification cycles and the amount of PCR product. Amplification was performed in a 50-µL reaction mixture using 5 µL (1:5 dilution) cDNA, platinum pfx Taq DNA polymerase (Invitrogen), and 1 µL deoxy-unspecified nucleoside 5'-triphosphate (dNTP) mixture (10 mM; Gibco-BRL), and specific MCP-1 primers recognize mouse cDNA at appropriate annealing temperatures. MCP-1 and GAPDH were amplified for 26 and 29 cycles, respectively. PCR products were separated on 2% agarose gel (NuSieve agarose, FMC, Rockland, ME). Primer sequences for MCP-1, VEGF-A, VEGF-B, VEGF-C, Ang-1, and GAPDH were described elsewhere [41
, 42
]. PCR primers for amplification of CCR2 and information on patterns of CCR2 tissue expression were from the Jackson Laboratory (Bar Harbor, ME; MGI:1205315, http://www.informatics.jax.org/). RNA from kidneys of C57BL/6 mice was used as a positive control. PCR amplification of CCR2 used 33 cycles. We adopted real-time quantitative RT-PCR (qRT-PCR) to evaluate the effect of PGE2 on the mRNA expression of VEGF-A, VEGF-B, VEGF-C, and Ang-1. qRT-PCR was performed using the Roche LightCycler system (Roche Molecular Biochemicals, Roche Diagnostics Co., Indianapolis, IN) with a SYBR-Green I dye (Molecular Probes, Junction City, OR). Amplification was performed in a 20-µL reaction mixture using 5 µL cDNA (1:5 diluted), 0.4 µL platinum Pfx Taq DNA polymerase (Gibco-BRL), 0.4 µL dNTP mixture (10 mM, Gibco-BRL), 0. 8 µl sense and antisense primer solution (50 µM), and SYBR-Green I dye at a 1:10,000 dilution. The sequences of primer pairs for VEGF-A, VEGF-B, VEGF-C, and Ang-1 were described previously [43
44
45
46
]. The optimal cycle program was determined for each gene in preliminary PCR runs to obtain a specific PCR product verified by melting curve analysis followed by gel electrophoresis. The absence of contaminating genomic DNA was ensured by RNA-PCR. The cDNA quantities were calculated by using the LightCycler analysis software, as described previously [47
]. VEGF-A, VEGF-B, VEGF-C, and Ang-1 were normalized to GAPDH [42
].
MC activation in vitro and detection of MC products
To evaluateVEGF-A production, MCs (106 cells/mL) were incubated in culture medium (RPMI 1640, Biofluids) containing 1% fetal calf serum (FCS; Biofluids) for 3 h and then stimulated by PGE2 (1000 nM) in complete medium. Ethanol (0.1%), used as a solvent of PGE2, was used as a relevant control for PGE2. After 12 h incubation, cell pellets were collected, and RNA was extracted for qRT-PCR. MC culture supernatants were obtained after cell culture for a period of 48, 72, 96, and 120 h and were used for VEGF-A detection by a specific enzyme-linked immunosorbent assay (ELISA; R&D Systems, Minneapolis, MN). The lower limit of VEGF-A detection was measured at 3 pg/mL. For time-course analysis of MCP-1 production and histamine release, MCs were washed twice with phosphate-buffered saline (PBS) and then incubated at 106 cells/mL for 0.5, 1, 2, 4, 6, or 24 h at 37°C in complete culture medium alone or with the addition of PGE2 (1000 nM). Ethanol (0.1%) was used as a relevant control for PGE2. To identify PGE2 receptor subtype(s) responsible for PGE2-induced MCP-1 secretion, BMMCs were incubated for 2 h with the following agents: PGE2 (1000 nM), butaprost (1000 nM), misoprostol (1000 nM), PGE1 alcohol (1000 nM), 17-phenyl--trinor-PGE2 (1000 nM), and sulprostone (1000 nM). Ethanol (0.1%) was used as a relevant control for these reagents. For quantification of intracellular MCP-1 under basal conditions, BMMCs were solubilized in lysis buffer (1% Nonidet P-40, 0.1% sodium dodecyl sulfate, 0.5% sodium deoxycholate in PBS) containing protease inhibitor cocktail (Boehringer Mannheim, Germany) plus sodium fluoride (20 mM), and cell debris was removed by centrifugation. MCP-1 content in the culture supernatant or cell lysates was measured by a specific ELISA (R&D Systems). The lower limit of detection for MCP-1 was measured at 2 pg/mL. At 0, 0.5, 1, 2, 3, 6, and 9 h time-points after incubation with PGE2 (1000 nM), cell pellets were collected as well, and RNA was extracted for semiquantitative RT-PCR analysis. To evaluate VEGF-A production, MCs (106 cells/mL) were incubated in culture medium (RPMI 1640 containing 1% FCS) for 3 h and then stimulated by PGE2 (1000 nM) in complete medium. Ethanol (0.1%) was used as a relevant control for PGE2. After 12 h incubation, cell pellets were collected, and RNA was extracted for qRT-PCR. MC culture supernatants were obtained after cell culture for a period of 48, 72, 96, and 120 h and were used for VEGF-A detection by a specific ELISA (R&D Systems). The lower limit of VEGF-A detection was measured at 3 pg/mL.
Histamine release measurement
To evaluate MC degranulation, we measured histamine release after PGE2 stimulation, as described previously [48
]. Briefly, MCs (106 cells/0.1 mL) were incubated in complete culture medium alone or with the addition of PMA (100 nM), calcium ionophore (1000 nM), and PGE2 at various concentrations (1–5000 nM) for 60 min at 37°C. Ethanol (0.1%) was used as a relevant control for these reagents. Residual histamine in the cell pellet was released by boiling the cells for 20 min, followed by spinning to remove cell debris. The concentration of histamine in the culture supernatant and pellet samples was measured by a specific ELISA, following the manufacturer’s instruction (Immuno-Biological Laboratories, Inc., Hamburg, Germany). The lowest limit of detection was determined at 2.4 ng/mL. Concentrations of MCP-1 in culture supernatants were measured by a specific ELISA described above. The net percent histamine release was calculated as follows: histamine in supernatant/(histamine in supernatant+histamine in pellet) x 100 (%).
Fluorescein-activated cell sorter (FACS) analysis
Cell-surface CCR2 was detected by flow cytometry as described previously [49
]. WEHI-3 cells, known to express low-level surface CCR2 [50
], were used as a positive control. Briefly, BMMCs, SPMCs, and WEHI-3 cells were incubated with 5 µg/ml monoclonal antibody (mAb) MC-21 (a kind gift of Dr. Matthias Mack, Klinikum University, Regensburg, Germany) or with an isotype control antibody (rat IgG2b, BD PharMingen, San Diego, CA) for 60 min on ice. After washings, the cells were incubated for 45 min on ice with an Alexa 488-labeled anti-rat polyclonal antibody (Molecular Probes; Invitrogen). After washing, CCR2 expression was assessed from 1.0 x 104 viable cells using a FACSCalibur cytofluorometer (Becton Dickinson, San Jose, CA) and analyzed using CELLQuest software (Becton Dickinson). Background fluorescence was assessed through staining with the isotype-matched antibody.
Ultrastructural localization of MCP-1
BMMCs maintained in culture as described above were fixed with 2.0% formaldehyde solution, followed by embedding in LR white plastic resin (EMBS Inc., Frederick, MD). Microthin sections (60–80 nm) were cut, mounted onto 200 mesh nickel grids, and processed for immunocytochemistry. Grids were incubated with 20 µg/ml hamster anti-mouse MCP-1 mAb: 2H5 (BD Pharminegen) or an isotype control mAb (BD PharMingen) for 4 h, followed by protein G conjugated to 10 nm gold particles (1/30, Polysciences, Warrington, PA). After 3 h incubation, sections were stained with uranyl acetate and Reynold’s lead. Sections were examined (EMBS Inc.) using a JEOL 1200 EX transmission electron microscope (JEOL Inc., Peabody, MA).
Studies of the effects of CsA and DM on MCP-1 secretion
We evaluated the effects of CsA, a cyclic peptide produced by the fungus Tolypocladium inflatum and the glucocorticoid DM on the release of MCP-1 from MCs under conditions previously reported to inhibit MC degranulation [51
, 52
]. MCs were pretreated with CsA at 1 µg/mL in complete culture medium for 1 h before addition of PGE2 (1000 nM). In some experiments, MCs were washed three times with PBS to remove CsA prior to addition of PGE2, as described previously for cytotoxic T lymphocytes [53
]. After 1 h incubation with PGE2, cell-free supernatants were tested for MCP-1 content. For DM treatment, MCs were preincubated with DM at 500 nM for 18 h and then incubated with PGE2, with or without washing, as described for CsA. Ethanol (0.1%), the solvent for CsA and DM, was used for a relevant control. All experiments were performed in triplicate. The percent MCP-1 release was calculated as follows: (MCP-1 release in CsA or DM group/MCP-1 release in control group) x 100 (%).
Measurement of MC proliferation
The effects of PGE2 on MC proliferation were assessed as described previously [54
]. Cells were washed twice with PBS, suspended in culture medium (RPMI containing 10% FBS and 10 ng/mL IL-3), plated (20,000 cells/well in 0.2 mL culture medium) in triplicate with the addition of PGE2 at various concentrations (1–5000 nM) onto 96-well plates, and incubated for 64 h. DNA synthesis was measured by [3H] thymidine deoxyribose ([3H]TdR) uptake (0.5 µCi/well, New England Nuclear, Boston, MA) during the last 16 h of culture. Radioactivity was measured by scintillation counting (Wallac Trilux Betaplate, Turku, Finland) after cell harvesting onto glass-fiber filtermates (Tomec Harvester 96 Mach III, Hamden, CT). The results are expressed as mean counts per minute (CPM; SD)/culture.
Statistical analysis
Statistical significance of group differences was evaluated by Student’s t-test using Excel software.
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800 pg/106 cells of intracellular MCP-1 under basal culture conditions, we found that PGE2 induces a rapid and massive release of MCP-1 from MCs (Fig. 1A
). After only 30 min exposure to PGE2, MC culture supernatants contained greater than 100 pg/ml MCP-1. Levels of MCP-1 reached a plateau of 700 pg/mL by 2 h (Fig. 1A)
and were maintained at approximately the same levels over a 5-day incubation period (not shown). Semiquantitative PCR analysis showed that BMMCs constitutively express MCP-1 mRNA, which is consistent with a previous report [55
], and that PGE2 minimally affects MCP-1 mRNA levels after 0.5, 1, 2, 3, 6, and 9 h incubation (Fig. 1B)
. PGE2 dose-dependently promoted MCP-1 release from BMMCs, and maximal effects were seen at PGE2 concentrations of 1000–5000 nM (Fig. 1C)
. Similar MCP-1 release was observed in SPMCs stimulated with PGE2: After 1 h stimulation with PGE2 at 1000 nM, MCP-1 levels in culture supernatant were 271.28 ± 9.90 pg/mL/106 cells as opposed to only 18.01 ± 0.68 pg/mL/106 cells in medium without PGE2. The viability of MC was not affected by short-term (2 h) and long-term (5 days) exposure to 1000 nM PGE2, as determined by trypan blue exclusion (data not shown).
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Figure 1. MCP-1 and histamine release from BMMCs induced by PGE2. (A) Time-course of MCP-1 and histamine release. BMMCs (106 cells/1 mL) were activated for a period of 0.5, 1, 2, 4, 6, and 24 h with PGE2 (1000 nM) or control (0.1% ethanol) solvent. MCP-1 and histamine were measured in the culture supernatant by specific ELISAs. Data shown reflect the mean (±SD) of triplicate determinations and are representative of five independent experiments. (B) Semiquantitative RT-PCR analysis of MCP-1 gene expression in BMMCs after exposure to PGE2 (1000 nM) or control 0.1% ethanol for the indicated time periods. RNA preparations were tested by parallel RT-PCR amplification for GAPDH. (C and D) Relationship between MCP-1 and histamine release from BMMCs (106 cells/0.1 mL), which were incubated with the addition of PMA (100 nM), the calcium ionophore A23187 (1000 nM), PGE2 at varying concentrations (1–5000 nM), or control (0.1% ethanol) for 60 min at 37°C. MCP-1 and histamine content in culture supernatants were measured by specific ELISAs. Histamine was also measured in the cell pellet after boiling for 20 min. The percent histamine release was calculated as follows: histamine in the supernatant/(histamine in the supernatant+histamine in the pellet) x 100 (%). The results shown reflect the means of triplicate determinations and were confirmed in an additional experiment. (E) RT-PCR analysis of CCR2 expression in BMMCs and SPMCs under basal culture conditions. RNA from mouse kidney served as a positive control. Results are representative of two independent experiments. (F) FACS analysis of CCR2 expression on BMMCs and WEHI-3 cells, which were cultured under basal conditions and were stained with a rat anti-mouse CCR2 mAb or isotype-matched control mAb, followed by an Alexa 488-labeled anti-rat IgG antibody. Results are representative of three independent experiments.
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RI receptor I or by thrombin and a variety of chemical mediators, is complete within 40 min [56
57
58
59
]. We assessed whether degranulation is the mechanism underlying MCP-1 release from MCs in the presence of PGE2. Using histamine release as a marker for MC degranulation [48
], we measured levels of MCP-1 (Fig. 1C)
and histamine (Fig. 1D)
in culture supernatants of MCs exposed to PGE2 (1–5000 nM) for 1 h. We found that PGE2 does not promote histamine release in BMMCs even at the highest concentration (5000 nM; Fig. 1D
), which reproducibly promotes MCP-1 release (Fig. 1C)
. By contrast, the calcium ionophore A23187, a strong stimulator of MC degranulation, induced histamine and MCP-1 release. In addition, PMA, which has been reported not to induce MC degranulation [60
, 61
], promoted MCP-1 but not histamine release from BMMCs (Fig. 1C)
. We also confirmed that PGE2 does not induce histamine release from MCs after longer incubation (Fig. 1A)
. In similar experiments, PGE2 and PMA promoted MCP-1 release from SPMCs (medium only: 18.01±0.68 pg/mL/106 cells; PGE2 5000 nM: 289.73±16.90 pg/mL/106 cells; PMA 100 nM: 298.36±9.92 pg/mL/106 cells at 1 h incubation) with minimal histamine release (medium only: 2.18±0.07%; PGE2 5000 nM: 17.30±0.23%; PMA 100 nM: 2.47±0.04%), whereas the calcium ionofore A23187 (1000 nM) induced histamine (72.51±2.28%) and MCP-1 (336.33±6.58 pg/mL/106 cells) release. We also found that SPMC released a minimal amount of histamine after 24 h exposure to PGE2 (medium only: 1.78±0.07%; PGE2 1000 nM: 5.70±0.70%). These results provide evidence that PGE2 can induce MCs to rapidly release MCP-1 without promoting their degranulation.
Analysis of CCR2 expression in MCs from mouse bone marrow and spleen
Previous studies revealed that MCP-1 is a potent inducer of degranulation from rat peritoneal MCs and murine pulmonary MCs but not from BMMCs [62
, 63
]. PGE2 was reported not to induce degranulation from BMMCs by itself but to facilitate degranulation induced by ionomycin or IgE/antigen [35
, 64
]. As shown above (Fig. 1A)
, we confirmed that PGE2 does not induce BMMC or SPMC degranulation alone or in conjunction with MCP-1 present in MC culture supernatant after incubation with PGE2. To investigate the reason for the failure of MCP-1 to promote degranulation in BMMCs and SPMCs, we evaluated expression of CCR2, the specific MCP-1 receptor. RT-PCR analysis revealed low-level CCR2 expression in BMMCs and SPMCs compared with a positive control RNA from mouse kidney (Fig. 1E)
, a result consistent with a previous report [65
]. These results were confirmed with MCs from a different passage (data not shown). By flow cytometry, CCR2 was undetectable on BMMCs (Fig. 1F)
and SPMCs (not shown). By contrast, surface CCR2 was detectable on WEHI-3 cells, which are known to express CCR2 only weakly [50
] (Fig. 1F)
.
Ultrastructural localization of MCP-1
Previous studies reported that BMMCs constitutively express MCP-1 mRNA at high levels [55
, 65
]. We found that murine MCs contain 800 pg MCP-1 protein/106 cells. The ability of PGE2 to induce the secretion of MCP-1 without histamine release (degranulation) in BMMCs suggested the involvement of a secretory process other than degranulation. Therefore, we examined the ultrastructural localization of MCP-1 in BMMCs maintained under basal culture conditions. As shown in Figure 2A
, BMMCs possess ultrastructural characteristics that are typical of MC, including a monolobed nucleus with partially condensed peripheral chromatin and numerous granules filled with electron-dense material. Immunochemical staining for MCP-1 revealed the presence of specific vesicle-like clusters of gold particles, which were not observed in control slides stained with an isotype-matched antibody (Fig. 2B
, arrowhead). In addition, MCP-1 was detected exclusively outside the MC-specific granules (Fig. 2B
, arrow). These results confirm the presence of pre-formed MCP-1 in BMMCs and provide evidence that MCP localizes outside the secretory granules in these cells.
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Figure 2. Cellular localization of MCP-1 in ultrathin sections of BMMCs detected by immunoelectron microscopy. (A) Representative image depicting the ultrastructure of BMMCs. (B) MCP-1 (arrowhead) detected in the cytoplasm of a BMMC by specific immunogold granule localization. The arrow points out a dense particle reflecting a secretory granule.
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Figure 3. Effects of CsA and DM on BMMC release of MCP-1. BMMCs (106 cells/mL) were pretreated with CsA (A) for 1 h or with DM (B) for 18 h; subsequently, PGE2 (1000 nM) was added directly into the culture or after cell-washing to remove residual CsA or DM. MCP-1, present in the culture supernatant, was measured by a specific ELISA. Ethanol (0.1%) was used as a control for PGE2. The percent MCP-1 released was calculated as follows: (MCP-1 release in CsA or DM group/MCP-1 release in control group) x 100 (%). Data shown reflect the means (±SD) of triplicate cultures and were confirmed in two repeat experiments. *, MCP-1 secretion from CsA- or DM-treated BMMCs is significantly different from that of control BMMCs (*, <0.05; **, <0.005).
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-trinor-PGE2 [68
] and sulprostone, which binds with high-affinity to the EP3 receptor and to a lesser extent, to the EP1 receptor [35
, 68
], induced the release of MCP-1 (Fig. 4B)
. Sulprostone induced greater MCP-1 release than 17-phenyl--trinor-PGE2 (P<0.05). The EP3/EP4 receptor-selective agonist PGE1 alcohol [68
] also induced MCP-1 release, whereas the EP2 receptor-selective agonist butaprost [36
, 68
] did not (Fig. 4B)
. Taken together, these results suggest that the PGE2 receptors EP1 and EP3 can mediate MCP-1 release from MCs, but involvement of the EP4 receptor cannot be ruled out.
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Figure 4. Effects of selective agonists of PGE2 receptors on MCP-1 release by BMMCs. (A) Listing of selective agonists for PGE2 receptors EP1, EP2, EP3, and EP4. High agonistic affinity toward a receptor subtype is noted (+); a lower affinity is noted (+/–). (B) BMMCs were treated with PGE2; with misoprostol (selective agonist for EP2/EP3 and EP4); with 17-phenyl--trinor-PGE2 (selective agonist for EP1); with sulprostone (selective agonist for EP1/EP3); with PGE1 alcohol (selective agonist for EP3/EP4); with butaprost (selective agonist for EP2); or with 0.1% ethanol (control) for 2 h. PGE2 and the EP agonists were used at a concentration of 1000 nM. MCP-1 released in the culture supernatant was measured by a specific ELISA. The results shown reflect the means (±SD) of triplicate cultures and are representative of three independent experiments.
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Figure 5. PGE2 modulation of VEGF-A, VEGF-B, VEGF-C, and Ang-1 expression in BMMC. (A) qRT-PCR analysis of gene expression in BMMCs treated for 12 h with PGE2 (1000 nM) or control 0.1% ethanol. Levels of VEGF-A, VEGF-B, VEGF-C, and Ang-1-specific mRNA, normalized to GAPDH, are expressed as fold increase above control. *VEGF-A induction from PGE2-treated BMMCs is significantly different from that of control-treated BMMC (*<0.05). (B) VEGF-A secretion by BMMCs treated with PGE2. BMMCs (106 cells/ml) were incubated in low serum medium (RPMI 1640 containing 1% FBS) for 3 h and then stimulated with PGE2 (1000 nM) in complete culture medium. Ethanol (0.1%) was used as a control for PGE2. VEGF-A was measured by a specific ELISA in BMMC culture supernatants obtained 48, 72, 96, and 120 h after PGE2 stimulation. The results reflect the means (±SD) of triplicate determinations and are representative of four independent experiments. (C) Effects of PGE2 on BMMC proliferation. Cells were washed twice with PBS, suspended in culture medium (RPMI containing 10% FBS and 10 ng/mL IL-3), plated (20,000 cells/well in 0.2 mL culture medium) in triplicate with various concentrations of PGE2 (1–5000 nM) onto 96-well plates, and incubated for 64 h. DNA synthesis was measured by [3H]TdR uptake (0.5 µCi/well) during the last 16 h of culture. Radioactivity was measured by scintillation counting after cell harvesting onto glass-fiber filtermats.
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Here, we report that PGE2 induces the prompt and massive release of the proangiogenic chemokine MCP-1 from murine MCs. This activity of PGE2 was not reported previously. Instead, it was noted previously that PGE2 inhibits MCP-1 release from synovial fibroblasts stimulated with IL-1ß [84
]. Recently, IL-1 was reported to induce MC release of newly synthesized IL-6, which was first detected 3 h after IL-1 exposure and peaked at 18 h [85
]. MCP-1 release from MCs is complete within 2 h from exposure to PGE2 and is not associated with increased MCP-1 gene transcription, suggesting that MCP-1 is secreted from pre-stored deposits. It is important that PGE2-induced MCP-1 secretion was not accompanied by degranulation. This observation is in agreement with previous studies showing that PGE2 alone does not induce MC degranulation [35
, 61
]. It is interesting that MCP-1 has been reported to induce degranulation from rat peritoneal MCs and murine pulmonary MCs but not from BMMCs [62
, 63
]. Here, we found that CCR2, the specific MCP-1 receptor, is undetectable on the surface of BMMCs and SPMCs, suggesting that insufficient levels of CCR2 are the likely cause for the failure of these cells to degranulate in response to MCP-1 induced by PGE2. Some studies have reported that PGE2 may even reduce MC degranulation induced by FcRI receptor I cross-linking with anti-IgE antibodies or by treatment with calcium ionophore A23187 [86
87
88
], but other studies have suggested that PGE2 can potentiate MC degranulation induced by ionomycin or by IgE/antigen complexes [35
, 64
]. Such differences could relate to heterogeneities among MC subtypes [89
]. We also found that PMA, which like PGE2, does not induce degranulation [35
], can promote MCP-1 release from MCs. Immunoelectron microscopy studies have provided evidence that IL-1-induced, newly synthesized IL-6 predominantly localizes in vesicle-like clusters, which are distinct from secretory granules, and is secreted from MCs independently of degranulation [85
]. Similarly, in the current study, immunoelectron microscopic analysis revealed that MCP-1 does not localize to the secretory granules in BMMCs but rather at distinct intracytoplasmic sites. Taken together, previous studies and the results presented here provide evidence that MCs can store synthesized cytokines at specific sites distinct from secretory granules and possess mechanisms for rapid exocytosis of pre-formed cytokines other than degranulation.
PGE2 exerts its activities on target cells through interaction with specific G protein-coupled receptors, of which there are four subtypes (EP1, EP2, EP3, and EP4). EP3 and EP4 receptors are ubiquitously expressed in tissues, whereas EP1 expression is somewhat more limited and is most abundant in the kidney [90 ]. By Northern analysis, BMMCs express EP1, EP3, and EP4 but not EP2 [61 ]. By RT-PCR analysis, low-level EP2 expression is detected in BMMCs [61 ]. By using selective receptor agonists to identify the EP receptors that mediate PGE2-induced MCP-1 secretion in BMMCs, we demonstrate that the PGE2 receptors EP1 and EP3 can mediate MCP-1 release from MCs, but a contribution of EP4 cannot be ruled out.
Several studies aimed at clarifying the proangiogenic properties of PGE2 have focused on its ability to induce VEGF-A secretion in different cell types [82 , 91 92 93 94 95 ]. Recently, it was noted that human MCs derived from the umbilical cord blood mononuclear cells constitutively produce VEGF-A and that PGE2 is approximately threefold more effective than the IgE-anti-IgE complex at stimulating VEGF-A release from IgE-sensitized MCs without causing degranulation [36 ]. The authors concluded that the EP2 subtype receptor is the main mediator of PGE2-induced VEGF-A secretion by human cord blood-derived MCs [36 ]. We could confirm that PGE2 promotes VEGF-A production in murine MCs, but the levels of VEGF-A measured in the culture supernatant of PGE2-stimulated murine MCs are lower than those detected in the culture supernatants of PGE2-stimulated human cord blood-derived MCs. The lower level of VEGF-A produced by murine MCs as opposed to human cord blood-derived MCs could be attributed to differences in distribution of PGE2 receptors. Human cord blood-derived MCs express EP1, EP2, EP3, and EP4 subtype receptors [36 ], and murine BMMCs express EP1, EP3, and EP4 but lack expression or express EP2 at low levels [35 , 61 ].
MCP-1 exerts its biological activities by binding and activating the G protein-coupled receptor CCR2 expressed on monocytes, endothelial cells, T lymphocytes, and natural killer cells. The release of MCP-1 by MCs induced by PGE2 is expected to recruit all these cell populations to sites of inflammation, as MCs have been reported to be a major source of MCP-1 [16
]. In particular, monocytes are a rich source of myriad angiogenic factors including VEGF-A, Ang-1, bFGF, transforming growth factor-ß1, platelet-derived growth factor, TNF-, hepatocyte growth factor, and insulin-like growth factor, suggesting that MCP-1 may exert some of its proangiogenic activities acting indirectly through the monocyte [23
, 79
, 96
, 97
]. It is interesting that recent studies about PGE2 modulation of MCP-1 activities have described a more vigorous monocyte chemotactic response to MCP-1 in the presence of PGE2 [98
], suggesting that PGE2 may exert additional stimulatory activities on monocytes. Endothelial cells have been shown to express a functional CCR2 receptor and to be responsive to MCP-1 [21
]. In addition to promoting endothelial cell chemotaxis, MCP-1 has been reported to directly stimulate endothelial cell sprouting [21
], a process that would be markedly enhanced by VEGF-A produced by MCs. Thus, MCP-1 and VEGF-A released by MCs during inflammation are likely to serve important, complementary, proangiogenic functions.
In summary, PGE2 induces murine MCs to promptly release MCP-1 and to increase VEGF-A secretion. These observations provide evidence supporting a role for PGE2 as a regulator of angiogenesis, in addition to inflammation and immunity. Understanding the diverse activities of PGE2 is critical to the development of new therapies for targeting inflammation.
Received April 25, 2005; revised September 28, 2005; accepted October 3, 2005.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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