Journal of Leukocyte Biology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published online as doi:10.1189/jlb.0106055 on February 27, 2007

Published online before print February 27, 2007
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
jlb.0106055v1
81/6/1445    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
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
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Bertram, C. M.
Right arrow Articles by Fogel-Petrovic, M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Bertram, C. M.
Right arrow Articles by Fogel-Petrovic, M.
(Journal of Leukocyte Biology. 2007;81:1445-1454.)
© 2007 by Society for Leukocyte Biology

Expression of kinin B1 and B2 receptors in immature, monocyte-derived dendritic cells and bradykinin-mediated increase in intracellular Ca2+ and cell migration

Cornelia M. Bertram*, Svetlana Baltic*, Neil L. Misso*, Kanti D. Bhoola*, Paul S. Foster{dagger}, Philip J. Thompson* and Mirjana Fogel-Petrovic*,1

* Lung Institute of Western Australia and Centre for Asthma, Allergy and Respiratory Research, The University of Western Australia, Perth, Australia; and
{dagger} Centre for Asthma and Respiratory Diseases, School of Biomedical Sciences, University of Newcastle, Newcastle, NSW, Australia

1 Correspondence: Lung Institute of Western Australia, Ground Floor, E Block, Sir Charles Gairdner Hospital, Nedlands, WA 6009, Australia. E-mail: mirjanaf{at}aari.uwa.edu.au


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The kinins, bradykinin (BK) and Lys-des[Arg9]-BK, are important inflammatory mediators that act via two specific G protein-coupled kinins, B1 and B2 receptors (B2R). Kinins influence the activity of immune cells by stimulating the synthesis of cytokines, eicosanoids, and chemotactic factors. Whether human dendritic cells (DC) express kinin receptors and whether kinins influence DC function are unknown. Fluorescence immunocytochemistry and RT-PCR were used to demonstrate that immature human monocyte-derived DC (hMo-DC) constitutively expressed kinins B1R and B2R. Kinin receptor expression was induced on the 3rd and 4th days of culture during differentiation of hMo-DC from monocytes and was not dependent on the presence of IL-4 or GM-CSF. Although monocytes also expressed B2R mRNA, the protein was not detected. The kinin agonists BK and Lys-des[Arg9]-BK up-regulated the expression of their respective receptors. BK, acting via the B2R, increased intracellular Ca2+, as visualized by confocal microscopy using the fluorescent Ca2+ dye, Fluor-4 AM. Evaluation of migration in Trans-well chambers demonstrated significant enhancement by BK of migration of immature hMo-DC, which was B2R-dependent. However, kinins did not induce maturation of hMo-DC. The novel finding that kinin receptors are constitutively expressed in immature hMo-DC suggests that these receptors may be expressed in the absence of proinflammatory stimuli. BK, which increases the migration of immature hMo-DC in vitro, may play an important role in the migration of immature DC in noninflammatory conditions and may also be involved in the recruitment of immature DC to sites of inflammation.

Key Words: human dendritic cells • chemoattractant • Lys-des[Arg9]-bradykinin


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Dendritic cells (DC) are potent APC, which reside in an immature state in the epidermal layer of the skin, in the respiratory and gastrointestinal systems, and in interstitial regions of solid organs, where they exert a sentinel function for incoming antigens [1 ]. Following an encounter with antigen, DC undergo a process of maturation, which enhances their antigen-presenting capacity and promotes their migration to lymphoid organs [2 , 3 ], where they induce T cell responses as well as immune tolerance [1 , 3 ]. The migration of DC in steady-state, noninflammatory conditions and the recruitment of DC and their precursors to sites of inflammation in response to chemotactic stimuli are critical for an optimum immune response [3 , 4 ].

Migration of DC to inflammatory sites is mediated by chemokines and their receptors. Locally produced, inflammatory chemokines (e.g., CCL3, CCL5, CCL2, and CCL20) mediate the recruitment of immature DC to sites of inflammation, whereas CCL21 and CCL19 are required for the migration of maturing DC to draining lymph nodes [5 ]. However, PGs [6 ], Type I IFN [7 ], and norepinephrine [8 ], as well as inflammatory stimuli such as TNF-{alpha}, IL-1ß [2 ], and possibly kinins, may also play a role in DC migration.

Kinins, specifically bradykinin (BK) and its metabolite Lys-[desArg9]-BK, are biologically active peptides with an established role in inflammation [9 , 10 ]. The cellular effects of kinins are mediated by highly specific, G protein-coupled receptors belonging to the seven transmembrane-spanning receptor family, which also includes a number of chemokine receptors [11 ]. The constitutively expressed B2 receptor (B2R) is widely distributed and mediates most of the biological actions of BK [11 , 12 ]. The inducible B1R is expressed in sepsis, inflammation, and tissue injury [12 , 13 ], and its expression requires induction by stimuli such as Lys-des[Arg9]-BK [14 ], IL-1ß [15 ], TNF-{alpha} [15 ], and infectious stimuli [16 ]. The human B2R is regulated at the protein level, and binding of BK results in rapid, receptor-mediated ligand internalization accompanied by loss of surface receptors [12 ]. However B1R, once induced under pathological conditions or stably expressed in culture, displays a lack of desensitization and absence of internalization [17 ]. Regulation of B1R expression by the B2R through activation of NF-{kappa}B has also been observed [18 ].

Multiple studies have confirmed the presence of kininogens, kallikreins, and kinin receptors in immune cells [13 , 19 20 21 22 ], supporting an important role for kinins in the immune response during inflammation. Furthermore, kinins strongly influence the activity of inflammatory cells by stimulating the synthesis of cytokines [20 ], eicosanoids [23 ], NO [24 ], and chemotactic factors [25 ]. B1R and B2R also play important roles in leukocyte migration [19 , 26 ]. However, the role of kinin receptors in human immature and mature DC is largely unknown, although one study in mice has demonstrated that kinins, acting through B2R, stimulate splenic DC to secrete IL-12 in vitro and in vivo [21 ]. In the current study, the hypothesis investigated was that human monocyte-derived DC (hMo-DC) express kinins B1R and B2R and that this expression is DC-specific. The effects of IL-4 and GM-CSF, cytokines necessary for the differentiation of monocytes into DC, on kinin receptor expression were also examined. In addition, the effects of BK and Lys-des[Arg9]-BK on the expression of their respective receptors and on intracellular Ca2+ DC cell migration and maturation were investigated.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Generation of hMo-DC
Human PBMC were isolated from heparinized blood by centrifugation on Ficoll-Paque (Amersham, Sydney, Australia) density gradients [27 ]. PBMC were resuspended in RPMI 1640 (Gibco-BRL, Grand Island, NY, USA) containing 10% heat-inactivated FCS (Life Technologies, Paisley, UK) and supplemented with glutamine (2 mM), gentamicin (60 ng/ml), penicillin (63 ng/ml), 50 µM ß-ME, and polymyxin B (1 µg/ml; Sigma Chemical Co., St. Louis, MO, USA), which were added to all culture media to negate any effects of trace amounts of endotoxin. Cells were allowed to adhere in 175 mm2 flasks. After 2 h, nonadherent cells were removed, and PBMC were cultured in RPMI 1640 containing recombinant GM-CSF (50 ng/ml) and IL-4 (10 ng/ml; R&D Systems, Minneapolis, MN, USA). Every 2–3 days, fresh culture media were added. By Day 7, 95–98% of cells were CD1a-positive, had a typical DC morphology, and expressed moderate levels of HLA-DR but little CD80 or CD86, as determined by flow cytometry. Less than 2% of cells in the culture were residual monocytes, as judged by expression of CD14.

Culture and stimulation of hMo-DC
To assess the effects of GM-CSF and IL-4 on B1R and B2R expression, cells were cultured in regular medium (complete RPMI 1640 containing GM-CSF and IL-4) for the first 6 days. On Day 6, cells were washed and placed in fresh medium containing GM-CSF only, IL-4 only, or neither of these cytokines and cultured for an additional 24 h. On Day 7, hMo-DC were stimulated with BK or Lys-des[Arg9]-BK for an additional 24 h, and the effects on B1R and B2R expression were assessed. BK and Lys-des[Arg9]-BK were used at a concentration of 10 µM, unless stated otherwise.

RT-PCR
Total RNA was isolated (RNeasy kit, Qiagen, Valencia, CA, USA) and DNase I-treated (Qiagen), and one-step RT-PCR (Qiagen) was performed. Specific sequences for sense (s) and antisense (as) primers for B1R and B2R were kindly provided by Dr. Carlos Figueroa (Institute of Histology and Pathology, Austral University of Chile, Valdivia, Chile). cDNA fragments of 213 bp for B1R mRNA (s: 5'-TTCTTATTCCAGGTGCAAGCAG-3'; as: 5'-CTTTCCTATGGGATGAAGATAT-3') and 335 bp for B2R mRNA (s: 5'-TGCTGCTGCTATTCATCATC-3'; as: 5'-CCAGTCCTGCAGTTTGTGAA-3') were amplified. The cDNA amplified from 18S rRNA (s: 5'-CATGCTAACTAGTTACG CGACC-3'; as: 5'-GAGCAAT AACAGGTCTGTGATG-3') was used as an internal control. The thermal cycler conditions were: cDNA synthesis at 50°C for 30 min and 95°C for 15 min (one cycle), followed by 94°C for 30 s, 52.4°C for 30 s, 72°C for 1 min (35 cycles), and 72°C for 10 min (one cycle). PCR products were resolved on 1.8% agarose gels.

Immunocytochemistry and confocal laser-scanning microscopy
The hMo-DC were washed in PBS, air-dried on polylysine-coated slides, and fixed for 10 s with methanol/acetone (1:1). Cells were subsequently blocked with 10% human serum (Sigma Chemical Co.) and 10% goat serum (Sigma Chemical Co.) in HBSS + IgG-free 1% BSA (Sigma Chemical Co.) and with protein blocker (DakoCytomation, Carpinteria, CA, USA) for 15 min. Primary rabbit antihuman antibodies to B1R or B2R (donated by Prof. Werner Müller-Esterl, Institute of Biochemistry II, University of Frankfurt, Germany) [28 29 30 ] were then applied for 2 h, and immunolabeling was performed using streptavidin-biotin + HRP (DakoCytomation) with 3-amino-9-ethylcarbazole (AEC; DakoCytomation) as the chromogen. For fluorescence labeling, receptor-bound B1R or B2R antibodies were linked to an AlexaFluor-conjugated goat antirabbit secondary antibody (Invitrogen, Mount Waverley, VIC, Australia) for 40 min in the dark. Excess secondary antibody was washed off, and preparations were mounted with Immu-mountTM (Thermo Shandon, Pittsburgh, PA, USA). Negative controls included hMo-DC labeled with nonspecific, primary rabbit polyclonal IgG (Dako) + AlexaFluor-conjugated goat anti-rabbit secondary antibody and with secondary antibody alone.

The HRP/AEC-stained slides were viewed under a Zeiss light microscope, whereas fluorescence-labeled slides were viewed on a BioRad MRC 1000/1024 UV laser-scanning confocal microscope at x600 magnification. Fluorescent images were artificially colored according to specific pixel brightness using a red-to-blue spectrum. For quantitative analysis, images were captured using an interference filter and inverted using ImagePro software. A circular area of interest (AOI) of constant size was drawn around each cell. The brightness of each pixel in the AOI was assessed and integrated, providing a fluorescence intensity value for the AOI. The integrated brightness of the AOI for the negative control was subtracted from the positively labeled slides. The resulting value represents the net signal intensity derived from specific immunolabeling. For statistical analysis, the intensities of 100 cells on each slide were averaged.

Calcium measurement: live cell imaging
DC cultured for 7 days were resuspended in RPMI without phenol red, containing 25 mM HEPES. For imaging by confocal microscopy, cells were allowed to settle on polylysine-coated coverslips for 1 h. The coverslips were placed in chambers (kindly provided by Dr. Paul Rigby, Biomedical Imaging and Analysis Facility, University of Western Australia, Perth) and incubated in RPMI without phenol red, containing 1% Pluronic F-127 and 2 µM Fluor-4 AM (Invitrogen) for 25 min at 37°C. Excess Fluor-4 AM was washed off, the chambers were placed under the confocal microscope, and 10 µM BK or 10 µM Lys-des[Arg9]-BK was added to the cells. The mean fluorescence intensity of six to eight cells in the visual field was measured in real time. Positive controls were mouse fibroblasts, which are known to produce a Ca2+ signal upon stimulation with BK [31 ], and hMo-DC stimulated to produce a Ca2+ signal with 1 µM fMLP (Sigma Chemical Co.) [32 ].

hMo-DC migration assay
Migration assays were performed in Trans-well chambers (Corning Inc., Hallam, VIC, Australia) with 1 x 106 cells/well. DC were incubated for 3 h in the upper chamber, which had 5 µm pores. To the lower chamber BK (10 µM), Lys-des[Arg9]-BK (10 µM), FCS (10%), or CCL5 (RANTES; 100 ng/ml; R&D Systems) was added. FCS [33 ] and in some of the experiments, CCL5, a chemotactic stimulus for immature DC [34 , 35 ], were used as positive controls. In experiments with the specific B2R antagonist, HOE-140, hMo-DC were first exposed to HOE-140 (1 or 10 µM; Sigma Chemical Co.) for 30 min, and BK (10 µM) was then added to the lower chamber. In some experiments, the B1R antagonist, Lys-[desArg9]-Leu8-BK (LDALB; 10 nM; Sigma Chemical Co.), was used together with HOE-140. Migrating cells were stained with trypan blue or vital dye (VibrantTM, Molecular Probes, Eugene, OR, USA). The number of migrating cells was evaluated by light or fluorescence microscopy. The results were expressed as a migration index (cells migrating in response to BK, Lys-des[Arg9]-BK, or FCS divided by cells migrating in response to medium alone). Chemokinesis was assessed by using the same concentration of BK (1 or 10 µM) in the upper chamber (with cells) and in the lower chamber. The effects of BK on DC locomotion were assessed after 4 h and expressed as the migration index.

Analysis of hMo-DC surface marker expression
Unstimulated cells and cells stimulated with BK ± HOE-140 or Lys-des[Arg9]-BK ± LDALB were washed in PBS supplemented with 10 mM NaN3 and resuspended in the same buffer containing 5% murine serum. Cells (105) were incubated for 30 min at 4°C with murine PE-conjugated anti-CD80, APC-conjugated anti-CD86, or PerCp-conjugated anti-HLA-DR (Becton Dickinson, San Diego, CA, USA). As a control, cells were stained with isotype-matched, irrelevant antibodies. After washing, cells were analyzed on a FACScan flow cytometer (Becton Dickinson).

Data analysis and statistics
For analysis of semiquantitative RT-PCR, gel-band intensities were determined using ImageQuant software (Molecular Dynamics, Sunnyvale, CA, USA) and normalized to the 18S rRNA bands. Intensity values were normally distributed, and therefore, mean (±SEM) intensity values were calculated, and differences between mean values were evaluated for statistical significance using Student’s t test.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Kinins B1R and B2R are expressed on DC
Analysis of B1R and B2R protein expression by fluorescence immunolabeling showed that immature, unstimulated hMo-DC expressed both receptors (Fig. 1a 1A and 1B ). However, monocytes did not express B1R or B2R protein (Fig. 1a 1D and 1E) . The specificities of the B1R and B2R antibodies were confirmed by use of nonspecific, primary rabbit polyclonal IgG antibodies (Fig. 1a 1C and 1F) . In hMo-DC, basal B2R mRNA expression was consistently greater than B1R mRNA expression, as determined by RT-PCR (Fig. 1b) .


Figure 1
View larger version (10K):
[in this window]
[in a new window]

  		Figure 1. hMo-DC express B1 and B2 kinin receptors. (a) Confocal microscopy images of differentiated, unstimulated hMo-DC on Day 7 of culture (A–C) and monocytes (hMo; D–F) after immunolabeling with kinin B1R (A and D) or B2R (B and E) primary antibodies and AlexaFluor-conjugated goat anti-rabbit secondary antibody. Nonspecific, primary rabbit polyclonal IgG (C and F) antibodies and AlexaFluor-conjugated goat anti-rabbit secondary antibody alone were used as negative controls (n=3). (b) B1R and B2R mRNA expression in hMo-DC as detected by RT-PCR (n=3); St, standard 100 bp ladder marker.

 
Expression of B1R and B2R is induced during differentiation of monocytes to DC
The differentiation of monocytes into DC is associated with functional and phenotypic changes. To investigate at what point during differentiation kinin receptor expression occurs, cells were collected on Day 1 of culture (monocytes) and on each of the following 7 days, and B1R and B2R expression was analyzed by immunolabeling (Fig. 2a and 2b ) and RT-PCR (Fig. 2d) . Monocyte cultures were assessed before the addition of GM-CSF and IL-4, which are required for differentiation of monocytes into hMo-DC. The absence of kinin receptor proteins on monocytes was confirmed. Expression of B1R and B2R was induced during in vitro differentiation, although the kinetics differed slightly with B1R, being first detected on Day 3, whereas B2R was first detected on Day 4 of culture (Fig. 2a and 2b) . B1R and B2R protein expression peaked on Day 5, which coincided with the full differentiation of monocytes to hMo-DC. The signal intensity per cell, as detected by fluorescence immunolabeling, increased on each day of culture for B1R and B2R (Fig. 2c) . Monocytes expressed B2R mRNA at a low basal level, whereas B1R mRNA was undetectable (Fig. 2d) . However, as B2R protein was not detected, it is possible that B2R synthesis is regulated post-transcriptionally. A significant increase in mRNA expression for both receptors was detected at Day 3 of culture. Figure 2e shows the fold increase in B1R and B2R mRNA expression relative to control during the 7 days of culture.


Figure 2
View larger version (49K):
[in this window]
[in a new window]

  		Figure 2. B1R and B2R protein and mRNA expression in monocytes and during differentiation of hMo-DC. Cells were collected on Day 1 (monocytes) and on Days 2–7 of culture during differentiation of hMo-DC. Representative images of B1R (a) and B2R (b) expression (HRP/AEC immunolabeling) in hMo-DC from one donor are shown. (c) Average signal intensity of AEC staining (red color) for B1R and B2R protein expression, recorded on 50 cells from four donors, for each day of culture. (d) Representative RT-PCR analysis of B1R and B2R mRNA expression and (e) densitometric values for B1R and B2R PCR products, relative to 18 S rRNA (internal control), on each day of culture (B1 mRNA, n=3; B2 mRNA, n=4).

 
Effects of GM-CSF and IL-4 on expression of B1R and B2R
Under physiological conditions, most cells do not express the inducible kinin B1R. Therefore, we investigated whether IL-4 or GM-CSF, which are required for differentiation of monocytes to DC, influenced the expression of kinin receptors. Cells cultured in complete medium (with IL-4 and GM-CSF) were used as a positive control. hMo-DC cultured with GM-CSF alone did not show any difference in B1R or B2R expression (Fig. 3a 3B and 3F ), whereas cells cultured with IL-4 alone showed a small, insignificant decrease in B1R and B2R expression (Fig. 3a 3C and 3G) , compared with hMo-DC cultured with GM-CSF and IL-4 (Fig. 3a 3A and 3E) . This suggested that B1R and B2R expression in unstimulated hMo-DC may be GM-CSF-dependent. In the absence of both cytokines, more pronounced decreases in B1R and B2R expression were observed, but these cells also showed changes in morphology and decreased viability (Fig. 3a 3D and 3H and 3b) .


Figure 3
View larger version (26K):
[in this window]
[in a new window]

  		Figure 3. Effects of GM-CSF and IL-4 on expression of B1R and B2R. (a) Representative images of B1R (A–D) and B2R (E–H) expression in hMo-DC (n=3) cultured for 7 days in GM-CSF + IL-4 (A and E) and in cells, which on Day 6, were transferred for an additional 24 h of culture in medium with GM-CSF alone (B and F), IL-4 alone (C and G), or without either cytokine (D and H). (b) The intensity of AEC staining (red color) for B1R and B2R expression was recorded on 50 cells from three donors for each of the different culture conditions and is presented as mean + SEM. There was a statistically significant decrease in B1R and B2R expression in hMo-DC cultured for 24 h without GM-CSF and IL-4 (*, P<0.05; n=4).

 
Effects of BK and Lys-des[Arg9]-BK on B1R and B2R mRNA and protein expression
As hMo-DC express B1R and B2R, we next examined the effects of the kinin receptor agonists, BK and Lys-des[Arg9]-BK, on the expression of B1R and B2R protein and mRNA. Stimulation with 10 µM BK or 10 µM Lys-des[Arg9]-BK increased the expression of B1R and B2R mRNA and protein in a time-dependent maner. Stimulation with a lower concentration of kinins (1 µM) up-regulates receptor expression to a similar degree (data not shown). Fluorescent labeling demonstrated a small, initial increase in B1R protein (between 15 min and 3 h) and a continued increase up to 6 h after stimulation with Lys-des[Arg9]-BK. The intense staining for B1R at 6 h (Fig. 4a ) was unchanged at 12 h poststimulation (data not shown). Similarly, B1R mRNA expression increased 15 min after stimulation with Lys-des[Arg9]-BK, suggesting de novo gene transcription and synthesis of B1R protein (Fig. 4a) . B2R protein peaked between 1 and 3 h, followed by a small decline at 6 h poststimulation (Fig. 4b) , indicating differential regulation of these two receptors at the protein level. After an initial, rapid increase in B2R mRNA expression 15 min after stimulation with BK, B2R mRNA levels continued to increase for at least 2 h and then declined (Fig. 4b) .


Figure 4
View larger version (35K):
[in this window]
[in a new window]

  		Figure 4. Effects of BK and Lys-des[Arg9]-BK on B1R and B2R protein and mRNA expression. Following stimulation with BK (10 µM) or Lys-des[Arg9]-BK (daBK; 10 µM) for 15 min–6 h, kinin receptors were detected by fluorescence immunolabeling using antibodies against B1R (a) or B2R (b; n=2). Color scale indicates pixel intensity. B1R and B2R mRNA expression 15 min–6 h after stimulation with Lys-des[Arg9]-BK (des-arg9 BK; a) or BK (b) was assessed by RT-PCR (n=3). The densitometric values (below images) indicate the fold increase in B1R and B2R mRNA in stimulated, relative to unstimulated, cells after normalization to 18S rRNA. Con, Control.

 
BK signaling in hMo-DC
An increase in intracellular calcium is an essential step in cell activation [31 , 36 ], and therefore, the effects of BK and Lys-des[Arg9]-BK on calcium flux in hMo-DC were assessed. Cells were loaded with the fluorescent Ca2+ dye Fluor-4 AM and then stimulated with the kinins. Exposure of hMo-DC to BK produced an increase in intracellular-free Ca2+ concentration (Fig. 5a ), whereas Lys-des[Arg9]-BK had no effect on intracellular Ca2+ (data not shown). In hMo-DC stimulated with BK, the increase in intracellular Ca2+ was much smaller than that induced by fMLP (Fig. 5a) . Thus, in addition to its effect on neutrophils and monocytes [32 ], fMLP is a potent stimulus for Ca2+ flux in hMo-DC. These data indicate that the magnitude of the Ca2+ flux in DC is dependent on the stimulus. In comparison with the signal obtained with BK in mouse fibroblasts, which were used as a positive control (Fig. 5b) , the signal in hMo-DC was smaller and of shorter duration.


Figure 5
View larger version (13K):
[in this window]
[in a new window]

  		Figure 5. Increase in intracellular-free Ca2+ concentration in hMo-DC following stimulation with BK. Average fluorescence emission from (a) eight live hMo-DC and (b) seven live mouse fibroblasts (Mouse F) in real time. Cells were loaded with 2 µM Fluor-4 AM, and (a) hMo-DC were stimulated with 10 µM BK and then with 1 µM fMLP (n=3), whereas (b) mouse fibroblasts (positive control) were stimulated with 10 µM BK and then with 1 µM thrombin. Signals from hMo-DC and mouse fibroblasts stimulated with BK were normalized to the maximum signals obtained with fMLP and thrombin, respectively (n=3).

 
Migration of hMo-DC in response to BK
The rationale for assessing the role of kinins in hMo-DC migration was that kinin receptors have been shown to play a role in leukocyte and neutrophil migration [26 ]. Furthermore, the expression of kinin receptors in immature hMo-DC, in the absence of proinflammatory stimuli, suggests that kinin receptors may have some function other than influencing the release of cytokines or DC maturation [19 ]. Therefore, kinins may also promote competency for migration in immature hMo-DC. There was a six- to tenfold increase in hMo-DC migration following stimulation with 10 µM BK (Fig. 6 ), whereas Lys-des[Arg9]-BK was ineffective in stimulating migration of immature hMo-DC. To confirm that the effect of BK on hMo-DC migration was mediated specifically by the B2R, cells were treated for 30 min with the B2R selective antagonist, HOE-140, prior to stimulation with BK. This resulted in decreased migration of immature hMo-DC, indicating that BK was acting via the B2R (Fig. 6) . Furthermore, when hMo-DC were pretreated simultaneously with a B2R antagonist (HOE-140) and a B1R antagonist (Lys-des[Arg9]-Leu8-BK), there was no additional inhibition of DC migration toward BK, when compared with HOE-140 alone, suggesting that the effect of BK was not mediated partially by the B1R (Fig. 6) .


Figure 6
View larger version (12K):
[in this window]
[in a new window]

  		Figure 6. Bradykinin activates migration of immature hMo-DC. (a) Migration assays were performed in Trans-well chambers with hMo-DC in medium + 1% FCS in the upper chamber. BK (10 µM), Lys-des[Arg9]-BK (daBK) (10 µM), HOE-140 (10 µM) + BK (10 µM) or HOE-140 (10 µM) + LDALB (10 µM) + BK (10 µM) in medium + 1% FCS were added to the lower chamber and incubated for 3 to 4 h. 10% FCS (n=9) and in some experiments, CCL5 (RANTES) (100 ng/ml) (n=3) were used as positive controls, whereas the negative control was medium + 1% FCS. Migrating cells were stained with trypan blue or vital dye (VibrantTM). The number of migrating cells was evaluated by light or fluorescence microscopy. Results are expressed as migration index (cells migrating in response to BK, Lys-des[Arg9]-BK, CCL5 or FCS divided by cells migrating in response to medium + 1% FCS). Each experiment (n=9) was performed in triplicate. (b) Chemotaxis of hMo-DC, with 1 or 10 µM bradykinin placed in the lower chamber only, compared with chemokinesis of hMo-DC, with the same concentrations of BK (1 or 10 µM) in both, the upper and lower chambers (n=3).

 
To distinguish whether BK only stimulated the motile apparatus of DC and the rate of locomotion (chemokinesis) or stimulated directed migration of DC toward the stimulus (chemotaxis), we compared the movement of DC when the same concentration of BK (1 or 10 µM) was used in both chambers, with the migration observed when BK (1 or 10 µM) was placed in the lower chamber only. The results indicate that relative to spontaneous locomotion (negative control), BK activated the cells and increased chemokinesis (Fig. 6b) . However, chemotaxis of DC toward 1 or 10 µM BK, placed only in the lower chamber, was much greater and more pronounced with 10 µM compared with 1 µM BK.

BK and Lys-des[Arg9]-BK do not induce phenotypic maturation of hMo-DC
By Day 7 of culture, immature hMo-DC expressed low levels of costimulatory (CD80, CD860) and MHC Class II (HLA-DR) molecules, as determined by flow cytometry (Fig. 7a , red line). As expected, the stimulation of hMo-DC with LPS for 24 h enhanced the expression of CD80, CD86, and HLA-DR (Fig. 7a , green line). The stimulation of immature hMo-DC with BK (Fig. 7b) or Lys-des[Arg9]-BK (Fig. 7c) for 24–48 h did not increase the expression of CD80, CD86, or HLA-DR in hMo-DC. B2R and B1R receptor antagonists (HOE-140 and LDALB, respectively) also did not alter CD80, CD86, or HLA-DR expression. Furthermore, there was no significant increase in IL-12 or PGE2 release in hMo-DC stimulated with kinins (data not shown).


Figure 7
View larger version (26K):
[in this window]
[in a new window]

  		Figure 7. Effects of BK and Lys-des[Arg9]-BK on CD80, CD86, and HLA-DR expression. Stimulated and unstimulated hMo-DC were labeled with CD80, CD86, or HLA-DR antibodies, and expression of CD80, CD86, and HLA-DR was analyzed by flow cytometry. Cells were stimulated for 24 h with BK (10 µM) or Lys-des[Arg9]-BK (10 µM), or cells were first exposed for 1 h to the B2R antagonist, HOE-140 (10 µM), or the B1R antagonist, LDALB (10 µM), and then stimulated with kinins for 24 h. (a) Histogram shows CD80, CD86, and HLA-DR expression in response to LPS (positive control; green line), relative to unstimulated DC (red line). (b) Expression of CD80, CD86, and HLA-DR in hMo-DC stimulated with BK (green line) or BK + HOE-140 (blue line) and (c) after stimulation with Lys-des[Arg9]-BK (daBK; green line) or Lys-des[Arg9]-BK + LDALB (blue line), relative to unstimulated DC (red line). Representative histograms from four independent experiments are shown.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
We report for the first time, that immature, unstimulated hMo-DC constitutively express the kinins B1R and B2R. The expression of B1R and B2R was detected at mRNA and protein levels. Constitutive expression of B2R has been reported in a number of other cell types, including vascular endothelial cells [37 ], human smooth muscle cells [38 ], murine and guinea pig macrophages [39 ], and mouse DC [21 ]. Although the inducible B1R is generally not expressed in normal tissues but is induced rapidly during inflammation [15 , 16 , 18 ], constitutive B1R expression has been reported in some species and cell types, including murine neutrophils [26 ], rat primary sensory A- and C-fibers [40 ], and macrophages [41 ]. In this study, constitutive expression of B1R was also detected in immature hMo-DC, suggesting that both kinin receptors may be expressed in the absence of proinflammatory stimuli.

Kinin receptor proteins were not detected in human monocytes, indicating that the expression of B1R and B2R is specific to hMo-DC. However, B2R mRNA was expressed at the basal level in monocytes, suggesting the possible presence on monocytes of low numbers of kinin B2R [42 ], which were below the detection limit of our methods. B1R and B2R mRNA expression during DC differentiation appeared to be differentially regulated, with the expression of B1R being detected 1 day earlier than B2R. Although in vivo studies point to the possibility that B1R may be up-regulated by B2R-mediated activation of NF-{kappa}B and autocrine production of proinflammatory cytokines [12 , 18 , 43 ], there is no information as to whether the opposite, namely, the up-regulation of constitutive B2R by activation of inducible B1R, may also occur. Moreover, as immature hMo-DC do not produce cytokines, except for low levels of IL-10, it is unlikely that proinflammatory cytokines are involved in autocrine regulation of B1R expression in hMo-DC.

As cytokines, such as IL-1ß, TNF-{alpha}, IL-8, and IL-4, can up-regulate expression of B1R [15 , 44 , 45 ] or B2R [46 ] in some cell types, the effects of IL-4 and GM-CSF on B1R and B2R expression were investigated. B1R and B2R expression did not differ in hMo-DC cultured for the last 24 h with GM-CSF alone or IL-4 alone, compared with cells cultured with both cytokines, suggesting that these cytokines are not critical for kinin receptor expression. Although more prolonged withdrawal of either cytokine may affect receptor expression, this cannot be tested, as such conditions affect DC morphology and viability. Thus, the significant decrease in B1R and B2R expression when hMo-DC were cultured without both cytokines may be a result of decreased cell viability.

BK, the primary endogenous agonist for B2R, and Lys-des[Arg9]-BK, the preferred agonist for B1R, transiently increased the expression of B2R and B1R mRNA, respectively, in hMo-DC. Although the B2R is usually constitutively expressed, up-regulation of B2R mRNA has been reported in cultured synovial and smooth muscle cells stimulated with platelet-derived growth factor and IL-1 [46 ]. In hMo-DC, the transient increase in B2R mRNA was rapid, indicating that BK was more likely to have induced de novo gene transcription rather than mRNA stabilization. The transient up-regulation of B1R mRNA following stimulation of hMo-DC with Lys-des[Arg9]-BK is in agreement with the kinetics of B1R mRNA expression in the human embryonic cell line IMR-90, costimulated with IL-1 and Lys-des[Arg9]-BK [47 ].

The rapid induction of both receptor proteins at the cell membrane supports a previous observation that in rat-isolated cardiac tissue, ischemia induced expression of B1R within 30–50 min [14 ]. However, this rapid (30–60 min) increase of both receptors at the cell membrane of hMo-DC might also suggest the existence of presynthesized receptors. Thus, stimulation with agonists may first facilitate transport of presynthesized receptor from the cytoplasm to the cell membrane, which could be followed by de novo transcription and receptor synthesis. Moreover, the kinetics of B2R and B1R expression are in agreement with reported differences in the regulation of kinin receptor proteins in other cell types. Thus, the decrease in B2R protein at 6 h may be a result of ligand-mediated internalization, accompanied by loss of cell surface receptors [31 ], whereas the continuous increase in B1R protein at the cell membrane may suggest a lack of desensitization and the absence of internalization [17 ].

The biological effects of kinins on human DC are largely unknown. The important roles of B1R and B2R in leukocyte migration [19 , 26 ], together with the findings that immature hMo-DC express B1R and B2R in the absence of proinflammatory stimuli, raised the possibility that kinin receptors are involved in the regulation of DC migration. In general, stimulation of the kinin B2R, which regulates the physiological actions of kinins, activates phospholipase C in a variety of cells types [48 ]. This results in the formation of diacylglycerol and inositol triphosphate, which stimulate the release of intracellular Ca2+ [38 , 49 ], an essential step in cell activation. In hMo-DC, the increase in intracellular Ca2+ seems to be associated with cell migration. In support of this, stimulation with Lys-des[Arg9]-BK did not increase intracellular Ca2+ and was also ineffective in inducing migration of hMo-DC. In contrast, BK was a potent stimulus for hMo-DC migration, relative to chemokinesis, indicating that BK is a chemoattractant for DC. The effect of BK was mediated by the kinin B2R, as migration of hMo-DC was inhibited by the B2R antagonist HOE-140. There was no additional inhibition of DC migration toward BK, when hMo-DC were pretreated simultaneously with a B2R antagonist and a B1R antagonist, indicating that the effect of BK was not partially mediated by the B1R. It seems that BK has differential effects on various immune cells and is an effective chemoattractant for immature hMo-DC but induces only moderate migration of human neutrophils and has no effect on eosinophil chemotaxis in vitro [41 ].

The first step in the mobilization of DC requires signals that promote hMo-DC competence for migration. The factors, inducing hMo-DC competence, include well-characterized DC maturation factors such as IL-1ß, TNF-{alpha}, and LPS [50 ]. In addition to enhancing DC migration by stimulating a Ca2+ signal and up-regulating B2R expression, BK may promote the competence of hMo-DC for migration. However, our findings that the expression of the costimulatory molecules, CD86 and CD80, and the capacity to present antigen (HLA-DR) were unchanged in hMo-DC stimulated with BK or Lys-des[Arg9]-BK suggest that kinins are not involved in phenotypic maturation of hMo-DC. In support of this, we did not observe an increase in IL-12 or PGE2 release in hMo-DC stimulated with kinins. Thus, in contrast to maturation factors that induce hMo-DC competence for migration, kinins and transforming growth factor ß1 (M. Fogel-Petrovic, C. M. Bertram, S. Baltic, N. L. Misso, K. D. Bhoola and P. J. Thompson and [51 ]) are the signals that induce DC migration but not maturation. Moreover, BK may increase DC competence for migration, possibly by activating the DC motile apparatus through a pathway other than maturation.

The absence of an increase in IL-12 release from hMo-DC following stimulation with BK contrasts with previous observations in murine DC [21 ]. In that study, kinins were shown to act through B2R to initiate secretion of IL-12 from mouse spleen DC in vitro and in vivo [21 ]. The discrepancy in the results for in vitro IL-12 release may be explained by the difference in the type of DC (Mo-DC vs. spleen DC) and/or the species difference of the investigated DC. The behavior and function of DC isolated from different organs or originating from different species are known to vary. For example, human and mouse bone marrow-derived and blood-derived DC have been reported to respond to CC chemokines [34 , 35 ], whereas mouse spleen DC did not migrate in response to any CC chemokine [52 ].

This study has shown that immature hMo-DC express kinins B1R and B2R in the absence of proinflammatory stimuli. BK and Lys-des[Arg9]-BK regulated the expression of their respective receptors at mRNA and protein levels. In addition, BK, acting via the B2R, increased intracellular Ca2+ and stimulated the migration of immature hMo-DC. Although the importance of BK in the complex interplay of molecules, which determines the overall pattern of DC migration in vivo, remains to be established, the in vitro data from this study strongly suggest that BK plays an important role in the migration of immature DC. Thus, it is possible that BK, which is released in vivo at low levels by a variety of cell types in the absence of inflammation and at higher levels in inflamed tissue, may mediate the up-regulation of B2R, the recruitment and migration of immature DC in noninflammatory conditions, as well as the recruitment of immature DC to sites of inflammation.


   ACKNOWLEDGEMENTS
 
This work was supported by funding from the National Health and Medical Research Council of Australia. We thank Dr. Paul Rigby (Biomedical Imaging and Analysis Facility, supported by Lotterywest) for help in the analysis of fluorescence-labeled hMo-DC by confocal laser-scanning microscopy. The authors acknowledge the assistance of the Red Cross of Western Australia in this research.

Received January 24, 2006; revised January 10, 2007; accepted January 11, 2007.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Banchereau, J., Steinman, R. M. (1998) Dendritic cells and the control of immunity Nature 392,245-252[CrossRef][Medline]
  2. Martin-Fontecha, A., Sebastiani, S., Hopken, U. E., Uguccioni, M., Lipp, M., Lanzavecchia, A., Sallusto, F. (2003) Regulation of dendritic cell migration to the draining lymph node: impact on T lymphocyte traffic and priming J. Exp. Med. 198,615-621[Abstract/Free Full Text]
  3. Sallusto, F., Lanzavecchia, A. (1999) Mobilizing dendritic cells for tolerance, priming, and chronic inflammation J. Exp. Med. 189,611-614[Free Full Text]
  4. Gunn, M. D., Kyuwa, S., Tam, C., Kakiuchi, T., Matsuzawa, A., Williams, L. T., Nakano, H. (1999) Mice lacking expression of secondary lymphoid organ chemokine have defects in lymphocyte homing and dendritic cell localization J. Exp. Med. 189,451-460[Abstract/Free Full Text]
  5. Sozzani, S., Allavena, P., D’Amico, G., Luini, W., Bianchi, G., Kataura, M., Imai, T., Yoshie, O., Bonecchi, R., Mantovani, A. (1998) Differential regulation of chemokine receptors during dendritic cell maturation: a model for their trafficking properties J. Immunol. 161,1083-1086[Abstract/Free Full Text]
  6. Kabashima, K., Sakata, D., Nagamachi, M., Miyachi, Y., Inaba, K., Narumiya, S. (2003) Prostaglandin E2-EP4 signaling initiates skin immune responses by promoting migration and maturation of Langerhans cells Nat. Med. 9,744-749[CrossRef][Medline]
  7. Asselin-Paturel, C., Brizard, G., Chemin, K., Boonstra, A., O’Garra, A., Vicari, A., Trinchieri, G. (2005) Type I interferon dependence of plasmacytoid dendritic cell activation and migration J. Exp. Med. 201,1157-1167[Abstract/Free Full Text]
  8. Maestroni, G. J. (2000) Dendritic cell migration controlled by {alpha} 1b-adrenergic receptors J. Immunol. 165,6743-6747[Abstract/Free Full Text]
  9. Cassim, B., Mody, G., Bhoola, K. (2002) Kallikrein cascade and cytokines in inflamed joints Pharmacol. Ther. 94,1-34[CrossRef][Medline]
  10. Bhoola, K. D., Figueroa, C. D., Worthy, K. (1992) Bioregulation of kinins: kallikreins, kininogens, and kininases Pharmacol. Rev. 44,1-80[Medline]
  11. Mahabeer, R., Bhoola, K. D. (2000) Kallikrein and kinin receptor genes Pharmacol. Ther. 88,77-89[CrossRef][Medline]
  12. Faussner, A., Bathon, J. M., Proud, D. (1999) Comparison of the responses of B1 and B2 kinin receptors to agonist stimulation Immunopharmacology 45,13-20[CrossRef][Medline]
  13. Calixto, J. B., Medeiros, R., Fernandes, E. S., Ferreira, J., Cabrini, D. A., Campos, M. M. (2004) Kinin B1 receptors: key G-protein-coupled receptors and their role in inflammatory and painful processes Br. J. Pharmacol. 143,803-818[CrossRef][Medline]
  14. Davis, A. J., Kelly, D., Perkins, M. N. (1994) The induction of des-Arg9-bradykinin-mediated hyperalgesia in the rat by inflammatory stimuli Braz. J. Med. Biol. Res. 27,1793-1802[Medline]
  15. Campos, M. M., Souza, G. E., Calixto, J. B. (1998) Modulation of kinin B1 but not B2 receptors-mediated rat paw edema by IL-1ß and TNF{alpha} Peptides 19,1269-1276[CrossRef][Medline]
  16. Passos, G. F., Fernandes, E. S., Campos, M. M., Araujo, J. G., Pesquero, J. L., Souza, G. E., Avellar, M. C., Teixeira, M. M., Calixto, J. B. (2004) Kinin B(1) receptor up-regulation after lipopolysaccharide administration: role of proinflammatory cytokines and neutrophil influx J. Immunol. 172,1839-1847[Abstract/Free Full Text]
  17. Zhou, X., Prado, G. N., Taylor, L., Yang, X., Polgar, P. (2000) Regulation of inducible bradykinin B1 receptor gene expression through absence of internalization and resensitization J. Cell. Biochem. 78,351-362[CrossRef][Medline]
  18. Phagoo, S. B., Poole, S., Leeb-Lundberg, L. M. (1999) Autoregulation of bradykinin receptors: agonists in the presence of interleukin-1ß shift the repertoire of receptor subtypes from B2 to B1 in human lung fibroblasts Mol. Pharmacol. 56,325-333[Abstract/Free Full Text]
  19. McFadden, R. G., Vickers, K. E. (1989) Bradykinin augments the in vitro migration of nonsensitized lymphocytes Clin. Invest. Med. 12,247-253[Medline]
  20. Tiffany, C. W., Burch, R. M. (1989) Bradykinin stimulates tumor necrosis factor and interleukin-1 release from macrophages FEBS Lett. 247,189-192[CrossRef][Medline]
  21. Aliberti, J., Viola, J. P., Vieira-de-Abreu, A., Bozza, P. T., Sher, A., Scharfstein, J. (2003) Cutting edge: bradykinin induces IL-12 production by dendritic cells: a danger signal that drives Th1 polarization J. Immunol. 170,5349-5353[Abstract/Free Full Text]
  22. Bhoola, K. D. (1996) Translocation of the neutrophil kinin moiety and changes in the regulation of kinin receptors in inflammation Immunopharmacology 33,247-256[CrossRef][Medline]
  23. Sakamoto, W., Kaga, M., Handa, H., Gotoh, K., Suzuki, S., Fujie, K., Inoue, N., Nishihira, J. (1994) Bradykinin and Met-T-kinin-Leu stimulated PGE2 production by rat macrophage and fibroblast Braz. J. Med. Biol. Res. 27,1923-1927[Medline]
  24. Bockmann, S., Paegelow, I. (1995) Bradykinin receptors in signal transduction pathways in peritoneal guinea pigs macrophages Eur. J. Pharmacol. 291,159-165[CrossRef][Medline]
  25. Sato, E., Koyama, S., Nomura, H., Kubo, K., Sekiguchi, M. (1996) Bradykinin stimulates alveolar macrophages to release neutrophil, monocyte, and eosinophil chemotactic activity J. Immunol. 157,3122-3129[Abstract]
  26. Araujo, R. C., Kettritz, R., Fichtner, I., Paiva, A. C., Pesquero, J. B., Bader, M. (2001) Altered neutrophil homeostasis in kinin B1 receptor-deficient mice Biol. Chem. 382,91-95[CrossRef][Medline]
  27. Romani, N., Reider, D., Heuer, M., Ebner, S., Kampgen, E., Eibl, B., Niederwieser, D., Schuler, G. (1996) Generation of mature dendritic cells from human blood. An improved method with special regard to clinical applicability J. Immunol. Methods 196,137-151[CrossRef][Medline]
  28. Haasemann, M., Buschko, J., Faussner, A., Roscher, A. A., Hoebeke, J., Burch, R., Muller-Esterl, W. (1992) Anti-idiotypic antibodies against the kinin receptor Agents Actions Suppl. 38,497-512[Medline]
  29. Abu Alla, S., Quitterer, U., Grigoriev, S., Maidhof, A., Haasemann, M., Jarnagin, K., Muller-Esterl, W. (1996) Extracellular domains of the bradykinin B2 receptor involved in ligand binding and agonist sensing defined by anti-peptide antibodies J. Biol. Chem. 271,1748-1755[Abstract/Free Full Text]
  30. Bhoola, R., Ramsaroop, R., Naidoo, S., Muller-Esterl, W., Bhoola, K. D. (1997) Kinin receptor status in normal and inflamed gastric mucosa Immunopharmacology 36,161-165[CrossRef][Medline]
  31. Ochsenbein, R. M., Inaebnit, S. P., Luethy, C. M., Wiesmann, U. N., Oetliker, O. H., Honegger, U. E. (1999) Differential regulation of bradykinin receptor density, intracellular Ca2+, and prostanoid release in skin and foreskin fibroblasts. Effects of cell density and interleukin-1{alpha} Br. J. Pharmacol. 127,583-589[CrossRef]
  32. Zhu, L., Castranova, V., He, P. (2005) fMLP-stimulated neutrophils increase endothelial [Ca2+]i and microvessel permeability in the absence of adhesion: role of reactive oxygen species Am. J. Physiol. Heart Circ. Physiol. 288,H1331-H1338[Abstract/Free Full Text]
  33. Corpataux, J. M., Naik, J., Porter, K. E., London, N. J. (2005) The effect of six different statins on the proliferation, migration, and invasion of human smooth muscle cells J. Surg. Res. 129,52-56[CrossRef][Medline]
  34. Vecchi, A., Massimiliano, L., Ramponi, S., Luini, W., Bernasconi, S., Bonecchi, R., Allavena, P., Parmentier, M., Mantovani, A., Sozzani, S. (1999) Differential responsiveness to constitutive vs. inducible chemokines of immature and mature mouse dendritic cells J. Leukoc. Biol. 66,489-494[Abstract]
  35. Dieu-Nosjean, M. C., Vicari, A., Lebecque, S., Caux, C. (1999) Regulation of dendritic cell trafficking: a process that involves the participation of selective chemokines J. Leukoc. Biol. 66,252-262[Abstract]
  36. Foyouzi-Youssefi, R., Petersson, F., Lew, D. P., Krause, K. H., Nusse, O. (1997) Chemoattractant-induced respiratory burst: increases in cytosolic Ca2+ concentrations are essential and synergize with a kinetically distinct second signal Biochem. J. 322,709-718[Medline]
  37. Smith, J. A., Webb, C., Holford, J., Burgess, G. M. (1995) Signal transduction pathways for B1 and B2 bradykinin receptors in bovine pulmonary artery endothelial cells Mol. Pharmacol. 47,525-534[Abstract]
  38. Schmidlin, F., Scherrer, D., Daeffler, L., Bertrand, C., Landry, Y., Gies, J. P. (1998) Interleukin-1ß induces bradykinin B2 receptor gene expression through a prostanoid cyclic AMP-dependent pathway in human bronchial smooth muscle cells Mol. Pharmacol. 53,1009-1015[Abstract/Free Full Text]
  39. Bockmann, S., Mohrdieck, K., Schmidt, H., Zundorf, G., Paegelow, I. (1998) Differential sensitivity of macrophages to bradykinin Naunyn Schmiedebergs Arch. Pharmacol. 357,151-158[CrossRef][Medline]
  40. Wotherspoon, G., Winter, J. (2000) Bradykinin B1 receptor is constitutively expressed in the rat sensory nervous system Neurosci. Lett. 294,175-178[CrossRef][Medline]
  41. Bockmann, S., Paegelow, I. (2000) Kinins and kinin receptors: importance for the activation of leukocytes J. Leukoc. Biol. 68,587-592[Abstract/Free Full Text]
  42. Rajasekariah, P., Warlow, R. S., Walls, R. S. (1997) High affinity bradykinin binding to human inflammatory cells Biochem. Mol. Biol. Int. 43,279-290[Medline]
  43. Pan, Z. K., Zuraw, B. L., Lung, C. C., Prossnitz, E. R., Browning, D. D., Ye, R. D. (1996) Bradykinin stimulates NF-{kappa}B activation and interleukin 1ß gene expression in cultured human fibroblasts J. Clin. Invest. 98,2042-2049[Medline]
  44. Bastian, S., Paquet, J. L., Robert, C., Cremers, B., Loillier, B., Larrivee, J. F., Bachvarov, D. R., Marceau, F., Pruneau, D. (1998) Interleukin 8 (IL-8) induces the expression of kinin B1 receptor in human lung fibroblasts Biochem. Biophys. Res. Commun. 253,750-755[CrossRef][Medline]
  45. Bryborn, M., Adner, M., Cardell, L. O. (2004) Interleukin-4 increases murine airway response to kinins, via up-regulation of bradykinin B1-receptors and altered signaling along mitogen-activated protein kinase pathways Clin. Exp. Allergy 34,1291-1298[CrossRef][Medline]
  46. Rehbock, J., Miska, K., Buchinger, P. (1999) Induction of the bradykinin B2-receptor, but not of the bradykinin B1-receptor, by interleukin-1ß in cultivated human decidua-derived cells Immunopharmacology 43,235-239[CrossRef][Medline]
  47. Menke, J. G., Borkowski, J. A., Bierilo, K. K., MacNeil, T., Derrick, A. W., Schneck, K. A., Ransom, R. W., Strader, C. D., Linemeyer, D. L., Hess, J. F. (1994) Expression cloning of a human B1 bradykinin receptor J. Biol. Chem. 269,21583-21586[Abstract/Free Full Text]
  48. Yano, K., Higashida, H., Inoue, R., Nozawa, Y. (1984) Bradykinin-induced rapid breakdown of phosphatidylinositol 4,5-bisphosphate in neuroblastoma X glioma hybrid NG108–15 cells J. Biol. Chem. 259,10201-10207[Abstract/Free Full Text]
  49. Llobet, A., Gual, A., Pales, J., Barraquer, R., Tobias, E., Nicolas, J. M. (1999) Bradykinin decreases outflow facility in perfused anterior segments and induces shape changes in passaged BTM cells in vitro Invest. Ophthalmol. Vis. Sci. 40,113-125[Abstract/Free Full Text]
  50. Cumberbatch, M., Dearman, R. J., Kimber, I. (1997) Interleukin 1 ß and the stimulation of Langerhans cell migration: comparisons with tumor necrosis factor {alpha} Arch. Dermatol. Res. 289,277-284[CrossRef][Medline]
  51. Sato, K., Kawasaki, H., Nagayama, H., Enomoto, M., Morimoto, C., Tadokoro, K., Juji, T., Takahashi, T. A. (2000) TGF-ß 1 reciprocally controls chemotaxis of human peripheral blood monocyte-derived dendritic cells via chemokine receptors J. Immunol. 164,2285-2295[Abstract/Free Full Text]
  52. Colvin, B. L., Morelli, A. E., Logar, A. J., Lau, A. H., Thomson, A. W. (2004) Comparative evaluation of CC chemokine-induced migration of murine CD8{alpha}+ and CD8{alpha}–dendritic cells and their in vivo trafficking J. Leukoc. Biol. 75,275-285[Abstract/Free Full Text]




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
jlb.0106055v1
81/6/1445    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
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
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Bertram, C. M.
Right arrow Articles by Fogel-Petrovic, M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Bertram, C. M.
Right arrow Articles by Fogel-Petrovic, M.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS