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Published online before print July 6, 2005
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* Department of Haematology, Barts and The London NHS Trust, United Kingdom;
Royal Veterinary College Royal College Street, London, United Kingdom; and
Diagnostic Oral Sciences, Clinical Sciences Research Centre, Queen Mary’s School of Medicine and Dentistry, London, United Kingdom
1Correspondence: Barts and The Royal London Hospital Medical School Haematology, Whitechapel, London, E1 1BB, UK. E-mail: marion.macey{at}bartsandthelondon.nhs.uk
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: protease-activated receptors • flow cytometry • cytokines • PAR-2
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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PAR-2 is expressed in a variety of tissues and is involved in cellular responses related to tissue injury and repair, angiogenesis, and perception of pain [1 , 3 4 5 ]. Recent research has shown the involvement of PAR-2 also in leukocyte function and inflammatory conditions. For example, human neutrophils express PAR-2 [6 , 7 ], and receptor gene knockout (PAR-2–/–) mice have significantly delayed inflammation, with reduced neutrophil margination and rolling on endothelium [8 ]. PAR-2–/– mice have further shown an important role for PAR-2 in type IV contact hypersensitivity reactions and chronic arthritis; they show distinctly diminished leukocyte infiltration and skin thickening 24 h and 48 h after challenge and remarkably, are completely resistant to adjuvant-induced arthritis [9 ]. The cellular mechanisms underlying these recent findings, however, are unknown.
The aim of the present study was to analyze whole blood from healthy volunteers for PAR-2 expression by flow cytometry. Of all mononuclear cells, only monocytes were consistently found to express cell-surface PAR-2. We therefore set out to investigate PAR-2 expression on purified monocytes and in vitro-differentiated macrophages and dendritic cells (DC) and also, the possibility that PAR-2 could act as a proinflammatory mediator for monocytes; the cells were examined for levels of intracellular calcium, interleukin (IL)-6, IL-8, IL-1ß, and tumor necrosis factor 
(TNF)- after receptor activation.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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, fetal liver tyrosine kinase 3 ligand (FLt3L), stem-cell factor (SCF), TGF-ß, IL-4, and interferon (IFN)-
were from R&D Systems (Abingdon, UK). Ficoll was from Amersham Biosciences Ltd. (Chalfont, St. Giles, UK). Fluo-3 was from Molecular Probes (Eugene, OR). PAR-2-activating peptide (AP; ASKH 95: 2-furoyl-LIGKV-OH) and control peptide (ASKH 115: phenyacetyl-LIGKV-OH) [9
, 10
] were prepared by Pheonix Pharmaceuticals (Belmont, CA). Before each set of experiments, peptides were reconstituted in media, placed in a waterbath sonicator for 1 min, vortexed, and then immediately added to the cells at the concentrations specified below. FluoroSpheres were from Dako (Ely, UK).
Antibodies
Purified, biotinylated, and fluorocein isothiocyanate (FITC)-conjugated mouse anti-human PAR-2 monoclonal antibody [mAb; immunoglobulin G2a (IgG2a), clone SAM-11], IgG2a isotype control, and goat anti-human PAR-2 polyclonal antisera N-19 were from Santa Cruz Biotechnology (CA). All other mAb and isotype controls used were from BD Biosciences (Oxford, UK) and include IL-1ß-phycoerythrin (PE; IgG1, AS10), IL-6-PE (IgG1, AS12), IL-8-FITC (IgG1, AS14), TNF--PE (IgG1, 6401.1111), CD68-PE (IgG2b, Y1/82A), CD14-PE or -FITC (IgG2a, M5E2), CD1a-FITC (IgG1, HI149), CD56-PE (IgG1, B159), CD3-PE (IgG1, UCHT1), CD19-PE (IgG1, HIB19). Streptavidin-FITC and FITC goat anti-mouse antiserum were from Southern Biotechnologies (Birmingham, AL). The horseradish peroxidase (HRP) rabbit anti-mouse serum for Western blotting was from Dako.
Cell lines
The human fibroblast cell line SVK14, promonocytic cell line U-937, and acute monocytic leukemia cell line THP-1 (American Type Culture Collection, Manassas, VA) were maintained in RPMI 1640 containing 1.5 mM L-glutamine and 10% FCS. All cell lines were mycoplasma-screened and confirmed negative.
Monocyte and CD34+ cell isolation
Venous blood from healthy adult volunteers was collected by venipuncture into heparinized tubes (Becton Dickinson, France). For CD34+ stem cell separation, leukapheresis samples from healthy adult volunteers were used. Peripheral blood mononuclear cells (PBMCs) were isolated by density gradient centrifugation. The magnetic antibody cell sorting (MACS) kit for negative selection of monocytes or the positive selection of CD34+ cells (Miltenyi Biotech, Bisley, UK) was used according to the manufacturer’s instructions and as described previously [11
] for the separation of monocytes and CD34+ cells, respectively. Samples of the resulting populations were immunostained to check the purity of the population; the monocyte population had less than 1% T cells, B cells, or natural killer (NK) cells, as confirmed by labeling with antibodies to CD3, CD19, and CD56, respectively. The CD34+ populations were at least 90% CD34+. In some experiments, monocytes were separated by plastic adhesion; PBMCs were plated at 5 x 106 cells/ml in RPMI 1640 containing 1.5 mM L-glutamine and 10% FCS (PAA Laboratories, Yeovil, UK) in six-well tissue-culture plates (Falcon, UK) and incubated at 37°C, 5% CO2, for 60 min. Nonadherent cells were washed away in prewarmed RPMI, and the adherent monocytes were gently collected using cell dissociation solution (Sigma Chemical Co.).
For inducing monocyte migration or cytokine production, cells were cultured at 1 x 106 cells/ml in M-SFM to which LPS (1 µg/ml), ASKH 95, or ASKH 115 (100 µM or as described below) was added. For cytokine production assays, brefeldin A was also added (5 µg/ml), and the cells were incubated at 37°C, 5% CO2, for 16 h before being analyzed for intracellular cytokine content by flow cytometry. For IL-1ß analysis, the absence of serum ensured intracellular storage of the protein [12 ].
Macrophage and DC differentiation
Purified monocytes were resuspended at a concentration of 2 x 106 cells/ml in RPMI 1640 or M-SFM containing 1.5 mM L-glutamine. For macrophage differentiation, M-SFM media were supplemented with 5% autologous serum. For DC differentiation, RPMI 1640 was supplemented with 10% FCS, 50 ng/ml GM-CSF, and 10 ng/ml IL-4. The cells were seeded in 24-well tissue-culture plates (Falcon) and incubated for 5 or 7 (DC) or 7 (macrophages) days in a humidified incubator at 37°C, 5% CO2. DC cultured for 7 days were given fresh media containing 50 ng/ml GM-CSF and 10 ng/ml IL-4 on Day 3 of culture. Purified CD34+ cells (95% CD34+, 0.5x106 cells/ml) were differentiated into DC by culture for 8 days (37°C, 5% CO2) in Cellgro media supplemented with 100 ng/ml GM-CSF and FLt3L, 10 ng/ml TNF-, and 20 ng/ml SCF.
Flow cytometry
For whole blood analysis, 60 µl blood anticoagulated with sodium citrate (0.105 M) was diluted 1:2 with fluorescein-activated cell sorter (FACS) buffer and incubated with 10 µg/ml relevant mAb on ice for 15 min. Red cells were lysed, and the samples were fixed using the Coulter TQ-prep system (Beckman Coulter, High Wycombe, UK) and then washed before analysis. Cultured cells were first incubated on ice for 20 min in FACS buffer [Ca- and Mg-free phosphate-buffered saline (PBS) containing 1% FCS, 1 mM EDTA, and 0.1% sodium azide] with purified mouse IgG2a (Dako) or normal mouse serum (1:200, Harlan Sera Lab, Loughborough, UK) to block nonspecific binding. For cell-surface labeling, 10 µg/ml relevant antibody was added, and samples were incubated for 20 min on ice, washed, resuspended in FACS buffer containing 5 µg/ml PI for dead cell exclusion, and immediately analyzed on a FACScan equipped with the CellQuest II software (Becton Dickinson, San Jose, CA). For intracellular labeling, cells were fixed in 3% formaldehyde, washed once in PBS, and then in FACS buffer containing 0.1% saponin prior to labeling with anticytokine or PAR-2/isotype mAb. After 20 min incubation on ice, samples were washed twice in saponin buffer and once in FACS buffer and analyzed on a FACScan. For data analysis, the CellQuest and WinMDI software was used. The relative fluorescence intensity for the expression of each cytokine was converted to molecules of equivalent soluble fluorochromes (MESFs) [13
] by use of a FluoroSpheres kit and following the manufacturer’s recommended procedure (Dako).
Calcium flux studies
To measure the changes in intracellular calcium in cells in response to PAR-2 AP and control peptide, the calcium indicator fluo-3 and flow cytometry were used. Cells (2x106ml–1) were incubated with fluo-3 at a final concentration of 2 µM for 30 min at 37°C in Ca- and Mg-free HBSS buffered with HEPES (1 mM). Cells were then diluted 1:10 in buffered HBSS containing Ca and Mg. The baseline level of fluo-3 fluorescence was determined in a dot plot of green fluorescence logarithmic scale on the y-axis and time in seconds on the x-axis. Then, the test or control peptide was added (100 µM final concentration), and the change in fluorescence was recorded. The calcium ionophore A23187 (1–2 µM final concentration) was used as a positive control. In some cases, monocytes, as present in PBMC preparations, were studied for PAR-2 responsiveness over time. For the restimulation experiments, a second and third dose of PAR-2 AP, each at a final concentration of 100 µM, was added at 5- or 20-min intervals. For the 20-min interval experiments, the fluo-3-labeled cells were incubated in Ca- and Mg-free HBSS buffered with HEPES (1 mM) in polypropylene tubes at 37°C 5% CO2 between stimulations. For all experiments, no washing or pipetting of the cells took place to minimize the possibility of mechanical-induced activation of the cells. Experiments were always ended by testing the response of the cells to the calcium ionophore to ensure that the prolonged incubation had not affected the capacity of the monocytes to generate a calcium flux.
Western blot analysis
SVK14 cells were brought into a single-cell suspension using a nonenzymatic cell dissociation solution. Granulocytes and PBMCs were isolated by density gradient centrifugation using Histopaque 1077 and 1119, according to the manufacturer’s recommendations. Monocytes were isolated as described above. The cells were lysed with nonionic detergent-containing buffer (radio immunoprecipitation assay), and the total protein was estimated by a bicinchoninic acid assay. The protein (5 µg per lane) was electropheresed on a 12% bis-acrylamide gel at 120 V and then transferred to a nitrocellulose membrane. After blocking with 5% milk overnight, the membrane was probed with mouse anti-PAR-2 antibody at room temperature (RT) for 2 h and then washed and visualized using a HRP-conjugated rabbit anti-mouse serum and enhanced chemiluminescence.
Statistical analysis
Significance of cytokine production and generation of calcium flux in restimulation experiments were calculated using two-way ANOVA.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The rapid up-regulation of cell-surface PAR-2 after mechanically induced stress suggested that the monocytes might have intracellular stores of PAR-2 protein. This was confirmed using intracellular labeling and flow cytometry (Fig. 2B)
. Furthermore, addition of brefeldin A to the blood before centrifugation blocked most of the up-regulation (Fig. 2B)
. Preincubation of whole blood with the PAR-2-activating or control peptide (100 µM), IFN- (100 ng/ml), TNF- (10 ng/ml), or LPS (1 µg/ml) for 2 h at RT or 37°C, however, did not increase surface PAR-2 expression (Fig. 2B
and data not shown). To the contrary, the storage of the whole blood at RT resulted in a complete loss of the discrete PAR-2 expression detected on the monocytes in freshly drawn blood (Fig. 2B)
. Moreover, monocytes separated from PBMCs by adhesion were found to lack surface PAR-2 expression (Fig. 2C)
.
The presence of PAR-2 protein in monocytes was confirmed by Western blotting, using two different goat polyclonal antisera and the SAM-11 mouse mAb (Fig. 3 ).
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PAR-2 activation results in calcium flux
Monocytes, freshly isolated by magnetic antibody sorting and monocyte-derived macrophages, produced a marked change in intracellular calcium when stimulated with the PAR-2 AP ASKH 95 (Fig. 5
, n=3). The use of whole PBMC preparations confirmed that monocytes and not lymphocytes responded to the PAR-2 AP (Fig. 5)
. The control peptide, ASKH 115, did not generate a calcium flux (Fig. 5) .
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, as determined by intracellular flow cytometry labeling (Fig. 7A
and data not shown). Dose responses showed that 12.5 µM ASKH 95 sufficed for IL-8 production over control peptide, and at least 25 µM was required for IL-6 and IL-1ß production (Fig. 7A)
. The control peptide induced some cytokine production over background production when tested at the highest doses only (Fig. 7A)
, and none of the peptides induced cytokine production in monocyte-derived DC (Fig. 7B)
.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The data presented here show that ex vivo peripheral blood monocytes have a discrete cell-surface expression of PAR-2 and similar to eosinophils [15
] and intestinal epithelial cells [16
], intracellular reservoirs of presynthesized PAR-2. The intracellular PAR-2 is apparently mobilized to the cell surface following centrifugation or repeated pipetting. Such a rapid up-regulation of PAR-2 in response to mechanical stress or injury is likely required for an appropriate response to the injury. Indeed, the finding that monocyte PAR-2 expression is sensitive to mechanical stress is supported by a study from Damiano et al. [17
], where PAR-2 expression in rat carotid artery was demonstrated to greatly increase after vascular balloon injury. These observations lead to the question of how the mobilization of intracellular PAR-2 is triggered. Few studies have addressed ways of increasing PAR-2 surface expression, but TNF- and LPS have been reported to up-regulate PAR-2 on epithelial cells after 20 h in vitro incubation [18
]. The PAR-2 up-regulation in the present report is much more rapid and unlikely to involve proinflammatory cytokines, as no influence on monocyte PAR-2 expression was detected after TNF-, IFN-, or LPS treatment of whole blood for up to 2 h. Another possibility is that factors released from activated or damaged cells are involved, for example, platelet-derived factors or PAR-2 agonists such as mast cell tryptase or neutrophil granzyme proteinase-3 [19
20
21
]. The latter possibility was investigated using ASKH 95, but this PAR-2 agonist alone did not affect monocyte PAR-2 expression. The involvement of PAR-2 agonist in PAR-2 up-regulation therefore seems less likely, and further research is required to identify factors involved in PAR-2 mobilization.
Contrary to the present study, Colognato et al. [22 ] recently reported human monocytes to be PAR-2-negative. It is important that we do not dispute the findings by Colognato et al. [22 ] but consider the results presented in their study and the present study in light of use of source and method for separation of monocytes. In their comprehensive study of protease-activated receptor expression on human monocytes, macrophages, and DC, Colognato et al. [22 ] mainly used monocytes separated from buffy coats by adherence. Although this source allows for a greater number of cells to be studied, it may have caused repeated activation of monocytes, during the process of buffy coat preparation and during the adhesion step [22 ]. It is well known that activation of monocytes can be caused by events such as adhesion to plastic surfaces [23 , 24 ]. In our control experiments, separation of monocytes from PBMCs by an adhesion step showed that such monocytes can be PAR-2-negative (Fig. 2C) . This highlights the importance of preparation procedures and its apparent effect on antigen expression, a subject we and others have previously addressed [25 ].
Repeated stimulation of PAR-2 has been shown to result in the desensitization of PAR-2 in epithelial cell lines and porcine endothelial cells [16 , 26 ]. In these studies, PAR-2 was internalized and degraded after receptor activation and resensitization that occurred through mobilization of intracellular stores and translation of existing mRNA but not by transcription [16 , 26 ]. Here, CD14+ monocytes were studied ex vivo in freshly drawn peripheral blood using whole blood flow cytometric analysis, and receptor activation studies were performed in whole blood or immediately after purification by MACS. Moreover, when the capacity of monocytes to generate calcium flux in response to repeated administration of the PAR-2 AP was investigated, it was found that a 20-min interval was required for the monocytes to show responsiveness again (Fig. 6) . After a further 40–60 min, however, the cells had no or only a weak response to the peptide, reflecting the down-regulation of receptor expression over time (Figs. 2 and 6) . It is therefore likely that the apparent divergent findings can be explained by the differences in cell source as well as technique used to separate monocytes.
In their recent commentary, Coughlin and Camerer [5 ] point to the possibility that PAR-2 could form part of a link between tissue injury and the decision to mount an adaptive immune response. The finding here that monocytes express functional PAR-2 receptors implicates these cells as candidates for forming such a link. It is interesting that tissue macrophages and DC in human skin have also been described as PAR-2+ [27 28 29 30 ], and both of these cell subsets can modify and in the case of DC, initiate adaptive immune responses. As monocytes can serve as precursors for macrophages and DC, the PAR-2 expression on monocyte-derived macrophages and DC was investigated. The monocyte-derived macrophages were found to be PAR-2-positive, and conversely, the majority of monocyte-derived DC lacked surface PAR-2, thus supporting the findings reported by Colognato et al. [22 ]. It should be mentioned that a small PAR-2+ subpopulation was observed consistently in the otherwise quite homogeneous DC population. These PAR-2-positive cells may represent a residual population of not fully differentiated monocytes, and it is possible that this would not be observed if a higher concentration of IL-4 had been used, as this cytokine has been shown to result in the down-regulation of surface PAR-2 expression [22 ]. Moreover, peripheral blood DC were found to be PAR-2-negative. The observation that DC precursors but not differentiated DC are PAR-2+ is in concordance with a recent study of DC development in mice, which demonstrated that PAR-2 was expressed on progenitor bone marrow cells but not bone marrow-derived DC or spleen-derived DC [31 ]. Of interest, we found that human acute myeloid leukemic blasts, which are classified on a morphological basis, differ in their expression of PAR-2, depending on differentiation stage; we did not detect PAR-2 on the immature myeloid precursor type blasts M1, M2, or M3, and the granulocytic and monocytic M4 and the monoblastic or monocytic M5-type blasts were PAR-2-positive (data not shown).
Monocytes are well described as major producers of a range of pro- and anti-inflammatory mediators, and the findings presented in this report raise the possibility that PAR-2 participates in the regulation of inflammatory responses through the induction of monocyte cytokine production. The production of IL-1ß, IL-6, IL-8, and TNF- by monocytes in response to the PAR-2 AP ASKH 95 was tested here. Consistent with previous reports for granulocytes, keratinocytes, gingival fibroblasts, and endothelial cells [21
, 32
, 33
], PAR-2-activated monocytes produced IL-6 and IL-8. The PAR-2-activated monocytes also produced IL-1ß, thus further suggesting a proinflammatory role for PAR-2. It has to be mentioned here, however, that IL-1ß- and IL-6-producing monocytes would also be expected to produce TNF-, and the apparent lack of TNF- production was an unexpected result. We must therefore stress the possibility that TNF- may have been produced but not detected by the assay used, and further research to establish a more complete picture of the cytokine production of monocytes in response to PAR-2 activation is planned.
PAR-2 is expressed at high density on motile cells such as granulocytes and tumor cells, and the capacity of certain tumor cells to migrate through collagen or fibronectin layers is increased following PAR-2 activation [34 , 35 ]. Leukocyte migration is also affected by PAR-2; topical administration of PAR-2 AP in rat postcapillary venules in vivo caused a significant increase in leukocyte rolling and adherence [36 ]. The possibility that human blood leukocytes would be stimulated to extravasate has not been addressed previously, and we therefore used human umbilical vein endothelial cells to test if PAR-2-activated monocytes would show increased transendothelial migration; however, this was not the case (data not shown). It should be noted that the PAR-2 peptide agonist has been reported as less able to induce cell migration compared with the physiological PAR-2 agonist tissue factor VIIa [35 ]. Indeed, it would be of great interest to study the response of monocytes stimulated with natural PAR-2 agonists; however, as several of these are known to or may mediate signaling via receptors other than PAR-2, the development of PAR-2-blocking antibodies or other antagonists would be useful for such studies.
In summary, this study shows for the first time that human monocytes are PAR-2+ and possess intracellular stores of PAR-2 protein. The data presented support the hypothesis that PAR-2 can contribute to proinflammatory immune responses, via monocyte production of IL-1ß, IL-6, and IL-8. Increased knowledge of how monocyte and macrophage PAR-2 expression is regulated in monocytes and macrophages and how these cells respond to PAR-2 activation is central to the further understanding of the pro- as well as potentially anti-inflammatory activity of PAR-2.
Received September 28, 2004; revised April 25, 2005; accepted May 9, 2005.
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B in human dermal microvascular endothelial cells J. Invest. Dermatol. 118,380-385[CrossRef][Medline]
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