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A New Staphylococcal Anti-Inflammatory Protein That Antagonizes the Formyl Peptide Receptor-Like 1¹

Cristina Prat^*,, Jovanka Bestebroer^*, Carla J. C. de Haas^*, Jos A. G. van Strijp^* and Kok P. M. van Kessel^2,*

^* Eijkman-Winkler Institute, University Medical Center Utrecht, Utrecht, The Netherlands; and Microbiology Department, Hospital Universitari Germans Trias i Pujol, Universitat Autonoma de Barcelona, Barcelona, Spain

	Abstract

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Bacteria have developed mechanisms to escape the first line of host defense, which is constituted by the recruitment of phagocytes to the sites of bacterial invasion. We previously described the chemotaxis inhibitory protein of Staphylococcus aureus, a protein that blocks the activation of neutrophils via the formyl peptide receptor (FPR) and C5aR. We now describe a new protein from S. aureus that impaired the neutrophil responses to FPR-like1 (FPRL1) agonists. FPRL1 inhibitory protein (FLIPr) inhibited the calcium mobilization in neutrophils stimulated with MMK-1, WKYMVM, prion-protein fragment PrP_106–126, and amyloid _1–42. Stimulation with low concentrations of fMLP was partly inhibited. Directed migration was also completely prevented toward MMK-1 and partly toward fMLP. Fluorescence-labeled FLIPr efficiently bound to neutrophils, monocytes, B cells, and NK cells. HEK293 cells transfected with human C5aR, FPR, FPRL1, and FPRL2 clearly showed that FLIPr directly bound to FPRL1 and, at higher concentrations, also to FPR but not to C5aR and FPRL2. FLIPr can reveal unknown inflammatory ligands crucial during S. aureus infections. As a novel described FPRL1 antagonist, it might lead to the development of therapeutic agents in FPRL1-mediated inflammatory components of diseases such as systemic amyloidosis, Alzheimer’s, and prion disease.

	Introduction

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Staphylococcus aureus remains a normal commensal of human skin and can potentially cause life-threatening infections involving any organ system (1). The ability of S. aureus to cause such a wide range of infections is the result of its extensive arsenal of virulence factors. Both bacterial surface components and secreted extracellular proteins have been described to contribute to the pathogenesis of infection. In addition, S. aureus uses efficient strategies to evade recognition by the innate immune system (1, 2).

Mobilization of phagocytes in response to chemoattractants constitutes the first line of defense against S. aureus infection. Chemoattractants are grouped in the superfamily of chemokines and the "classical" chemoattractants, which include the formylated peptides (side products of bacterial translation), activated complement component 5 (C5a) and C3 (C3a), leukotriene B₄ (LTB₄),³ and platelet-activating factor (PAF). Both classical chemoattractants and chemokines activate seven-transmembrane G protein-coupled receptors expressed on cells of hemopoietic origin but also on many other cell types (3, 4).

We recently described the chemotaxis inhibitory protein of S. aureus (CHIPS) (5), an excreted protein that impairs the response of neutrophils and monocytes to C5a and formylated peptides such as fMLP. CHIPS binds directly to the C5aR and formyl peptide receptor (FPR) preventing the natural ligands from activating these receptors (6).

FPR is the high-affinity receptor for fMLP that is activated by picomolar to nanomolar concentrations of fMLP and is expressed on phagocytes but also on cell types as diverse as hepatocytes, dendritic cells, astrocytes, and microglia cells (7, 8, 9). Two other homologs of the FPR have been identified, FPR-like 1 (FPRL1) and FPR-like 2 (FPRL2). Human phagocytes are known to differentially express these receptors. Neutrophils express the FPR and FPRL1, whereas monocytes and basophils express all three members FPR, FPRL1, and FPRL2 (10, 11, 12, 13). Mature dendritic cells express FPRL2, low levels of FPR, but no FPRL1 (12). FPRL1 is considered a low-affinity fMLPR and is expressed in an even greater variety of cell types. In the last years, a wide variety of agonists for this receptor have been identified, including components from microorganisms and host-derived peptide and lipid agonists (11, 14). It is remarkable that the FPRL1 is used by at least three amyloidogenic ligands: serum amyloid A (15), the 42-aa form of amyloid (A_1–42; Ref. 16), and the prion protein fragment PrP_106–126 (17). These ligands have been shown to attract phagocytes with important implications in pathological states such as systemic amyloidosis, Alzheimer’s disease (18), and prion disease, respectively. Several small synthetic peptides such as MMK-1 (19), WKYMVm (20), and WKYMVM (10, 21) were selected from random peptide libraries and have been identified as agonists for the FPRs and are widely used for research purposes. Recently, F2L, an acetylated peptide derived from the human heme-binding protein, was identified as a new natural chemoattractant agonist specific for FPRL2 (22).

The importance of CHIPS as a potential virulence factor led us to investigate homologous excreted proteins in the genome of S. aureus. A gene was found that showed 49% homology with the gene for CHIPS (chp) and contained a leader peptide and a peptidase cleavage site (amino acid sequence AXA). The gene encodes for a cleaved 105-aa protein with 28% homology with CHIPS. Initial functional assays with the recombinant protein demonstrated a weaker but consistent inhibition of fMLP-induced activation of neutrophils. Further analysis demonstrated that this new protein impairs the neutrophil and monocyte responses to FPRL-1 agonists.

In this study, we describe a new protein from S. aureus with anti-inflammatory properties, FPRL1 inhibitory protein (FLIPr). We show that FLIPr inhibits the leukocyte response to FPRL1 agonists and we demonstrate binding of FLIPr to HEK293 cells expressing the FPRL1.

	Materials and Methods

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Reagents

MMK-1 (LESIFRSLLFRVM) was synthesized by Sigma-Genosys. fMLP, rC5a, anti-FLAG mAb, propidium iodide, and L--lysophosphatidylcholine were obtained from Sigma-Aldrich. WKYMVm was synthesized by Dr. J. A. W. Kruijtzer (Department of Medicinal Chemistry, Utrecht Institute for Pharmaceutical Sciences, Utrecht, The Netherlands). WKYMVM, PrP_106–126, and amyloid peptide A_1–42 were obtained from Bachem. IL-8 and growth-related oncogene (GRO)- were purchased from PeproTech. PAF-16 was obtained from Calbiochem. LTB₄ was obtained from Cayman Chemical. Lipoxin A4 was obtained from BIOMOL. Fluo-3-AM (acetoxymethyl ester), calcein-AM, Fura red-AM, fura 2-AM, and Alexa Fluor 488 phalloidin were obtained from Molecular Probes. Anti-hemagglutinin (HA) mAb (clone 12CA5) was obtained from Roche Applied Science. Allophycocyanin-labeled goat anti-mouse Ig was obtained from BD Pharmingen. PE-conjugated mAbs CD4-PE (Leu-3a), CD8-PE (Leu-2a), CD19-PE (Leu-12), CD56-PE, CD16-PE, and CD14-PE (Leu-M3) were obtained from BD Biosciences; CD3-R-PE-Cy5 (clone UCHT1) was obtained from DakoCytomation.

DNA sequence

The program tblastn with the nonredundant DNA database and the S. aureus genome database at www.ncbi.nlm.nih.gov was used to check for sequence similarities with the chp gene. A gene was found with a 49% homology with chp. The DNA sequence of the gene encoded a protein of 105 aa (in bold), preceded by a signal peptide and a signal-peptidase site (underlined): MKKNITKTIIASTVIAAGLLTQTNDAKAFFSYEWKGLEIAKNLADQAKKDDERIDKLMKESDKNLTPYKAETVNDLYLI VKKLSQGDVKKAVVRIKDGGPRDYYTFDLTRPLEENRKNIKVVKNGEIDSIYWD.

Primers were designed according to the published sequence of the gene (hypothetical protein SAV1156, S. aureus subspecies aureus Mu50; gene ID 1121132) for the cloning of the protein into pRSET vector (Invitrogen Life Technologies) and were manufactured by Invitrogen Life Technologies.

Prevalence in clinical S. aureus isolates

Prevalence of the gene for FLIPr (flr) was checked in 91 clinical and laboratory S. aureus isolates. Genomic DNA was isolated from cultures of S. aureus using the High Pure PCR Template Preparation kit (Roche). PCR amplification was conducted using Supertaq polymerase (Enzyme Technologies) and 5'-TTCTTTAGTTATGAATGGAA-3' as the forward primer and 5'-TTAATCCCAATAAATCGAGTCG-3' as the reverse primer. PCR products were detected by electrophoresis through agarose gel and ethidium bromide staining.

Cloning and expression of the protein

The flr gene, without the signal sequence, was cloned into the pRSET vector directly downstream of the enterokinase cleavage site and in frame of the EcoRI restriction site by overlap extension PCR (23). The plasmid pRSET was used as template for amplification of DNA fragments having overlapping ends using the sense primer 5'-GCTCTAGAAATAATTTTGTTTAACTTTAAGAAGGAG-3' containing XbaI restriction site (underlined nucleotides) and the antisense primer 5'-TCTAAACCTTTCCATTCATAACTAAAGAACTTGTCGTCATCGTCGTACAG-3'. The gene was then amplified by PCR on chromosomal DNA of S. aureus Newman using the sense primer 5'-TTCTTTAGTTATGAATGGAA-3' and the antisense primer 5'-CGTCCTGAATTCTTAATCCCAATAAATCGAGTCG-3', containing the EcoRI restriction site (underlined nucleotides). The obtained DNA fragments were mixed, denatured, and reannealed in a subsequent PCR, using the primers corresponding to the 5' and 3' end sequences, to obtain the full-length PCR product. The amplification reactions were performed using PfuTurbo DNA polymerase (Stratagene). The final PCR product was purified using PCR Purification kit (Qiaquick; Qiagen), cloned into the EcoRI and XbaI site of the pRSET vector and propagated in TOP10F' Escherichia coli following manufacturer’s instructions (Invitrogen Life Technologies). After verification of the correct sequence by using ABI Prism 377 (Applied Biosystems), the recombinant protein was expressed in Rosetta-Gami E. coli (De3)pLysS (Novagen; Merck Biosciences) by induction with 1 mM isopropyl -D-thiogalactoside (Invitrogen Life Technologies).

Purification and FITC labeling of the protein

Bacteria were lysed with CelLytic B Bacterial Cell Lysis/Extraction Reagent (Sigma-Aldrich) and lysozyme according to the manufacturer’s description. The histidine-tagged protein was purified using a nickel column (HiTrap Chelating HP, 5 ml; Amersham Biosciences) following the manufacturer’s instructions and cleaved afterward with enterokinase (Invitrogen Life Technologies). Samples were checked for purity and presence of protein by means of 15% SDS-PAGE (Mini Protean III System; Bio-Rad) and Coomassie brilliant blue (Merck) staining.

A portion of the protein was labeled with FITC (Sigma-Aldrich) for binding experiments. Therefore, 500 µg/ml FLIPr was incubated with 50 µg/ml FITC in carbonate buffer (pH 9.0) for 1 h at 37°C under constant agitation. FLIPr-FITC was separated from unbound FITC using a desalting column (HiTrap desalting; Amersham Biosciences). The fractions were collected and tested for the presence of FLIPr (OD₂₈₀) and FITC (OD₄₉₅) in a spectrophotometer, to calculate the concentration: FLIPr – FITC (mg/ml) = (OD₂₈₀ – (0.35 x OD₄₉₅))/1.547. rCHIPS was isolated, purified, and FITC labeled as described (5) using essentially the same procedures as for FLIPr.

Leukocyte isolation

Venous blood was collected from healthy volunteers into tubes containing sodium heparin. Blood was diluted with an equal volume of PBS and layered onto a gradient of 12 ml of Histopaque (density 1.119; Sigma-Aldrich) and 10 ml of Ficoll (Amersham Biosciences) and centrifuged for 20 min at 380 x g and 21°C. PBMC and polymorphonuclear neutrophils (PMN) were collected separately from Ficoll and Histopaque interphases, respectively. Cells were then washed with cold RPMI 1640 (containing 25 mM HEPES and L-glutamine; BioWhittaker) with 0.05% human serum albumin (RPMI 1640-HSA). For elimination of erythrocytes, the PMN pellet was subjected to a hypotonic shock by adding ice-cold H₂O for 30 s and subsequently 10 times concentrated PBS to reconstitute isotonicity. Cells were finally resuspended to a concentration of 1 x 10⁷ cells/ml in RPMI 1640-HSA and used promptly.

Human embryonic kidney (HEK) 293 cells

HEK cells were transiently transfected with plasmids containing the DNA encoding a FLAG-tagged version of the human membrane receptors FPR, FPRL1, and C5aR or a 3xHA-tagged FPRL2. The DNA sequence of the receptors was amplified by PCR by using the following primer pairs: for FPR, sense primer 5'-CCGGAATTCATGGACTACAAGGACGACGACGACAAGATGATGGAGACAAATTCCTCTCTC-3' and antisense primer 5'-GCTCTAGATCACTTTGCCTGTAACGCCAC-3'; for FPRL1, sense primer 5'-CCGGAATTCATGGACTACAAGGACGACGACGACAAGATGGAAACCAACTTCTCCACTCCTC-3' and antisense primer 5'-GCTCTAGATCACATTGCCTGTAACTCAG-3'; for C5aR sense, primer 5'-CCGGAATTCATGGACTACAAGGACGACGACGAC AAGATGAACTCCTTCAATTATACC-3' and antisense primer 5'-GCTCTAGACTACACTGCCTGGGTCTTCT-3'.

Primers contained EcoRI and XbaI restriction sites (in bold nucleotides). An N-terminal FLAG-tag (DYKDDDDK, included in the sense primers, underlined nucleotides) was placed after the first methionine for detection by the anti-FLAG M2 mAb. An extra methionine (ATG) was included directly after the FLAG tag to keep the N-terminal sequence intact. The amplification reaction was performed on human bone marrow QUICK-Clone cDNA (BD Biosciences/BD Clontech) using PfuTurbo DNA polymerase. The PCR product was digested with EcoRI and XbaI, ligated in the expressing plasmid pcDNA3.1 (Invitrogen Life Technologies) and transfected into HEK293 cells as described previously (24). The 3xHA-tagged FPRL2 DNA was obtained from the University of Missouri-Rolla cDNA Resource Center (Rolla, MO) and was also transfected into HEK293 cells. HEK293 cells were grown in a 6-well plate (Costar) at 0.5 x 10⁵ cells/ml and maintained in Eagle’s MEM (BioWhittaker) supplemented with 0.1 mM nonessential amino acids, 1 mM sodium pyruvate, 10 µg/ml gentamicin, and 10% FCS. After 3–4 days of culture, cells were transfected with the respective plasmids by using Lipofectamine 2000 (Invitrogen Life Technologies), according to manufacturer’s instructions. After 2–3 days from transfection, cells were used for binding assays.

Calcium mobilization

The activation of neutrophils by chemoattractants initiates a rapid and transient increase in the free intracellular calcium concentration. Calcium mobilization with isolated human neutrophils and monocytes was measured as previously described (25). Briefly, the PMN fraction (5 x 10⁶ cells/ml) was loaded with 2 µM Fluo-3-AM or fura red-AM for 20 min at room temperature, protected from light, and kept under constant agitation. Cells were washed, resuspended in RPMI 1640-HSA, and incubated with buffer or protein (FLIPr or CHIPS) for 20 min. The cells (1 x 10⁶ cells/ml) were then monitored for calcium mobilization over time, first for 10 s to determine the basal fluorescence level, and then for 40 s after addition of the concentrated stimulus. Fluorescence was measured at 530 nm (for Fluo 3-AM) or 560 nm (for Fura red-AM) using a flow cytometer (FACSCalibur or FACScan; BD Biosciences). For calcium mobilization in PBMC, a PE-conjugated anti-CD14 was included during labeling with Fluo 2-AM. PBMC were adjusted to 5 x 10⁶ cells/ml and monocyte calcium mobilization was monitored by gating on side scatter and anti-CD14 staining. Alternatively, neutrophils were labeled with 2 µM fura 2-AM for 45 min at room temperature, washed, and resuspended in HBSS (BioWhittaker) containing 1% HSA at 7.5 x 10⁶ cells/ml. Cells were transferred into black clear-bottom microtiter plates (50 µl) and preincubated for 5 min with 25 µl of inhibitory protein (FLIPr or CHIPS) or HBSS-HSA buffer control and subsequently loaded into a FlexStation fluorescent plate reader (Molecular Devices). Fluorescence was measured every 1.5 s at dual emission wavelengths of 530 and 590 with 340 excitation. Stimuli (25 µl) were automatically added after a 1 min baseline reading and measurement continued for an additional 5 min. The ratio of 530:590 was calculated for every reading and plotted vs time.

Changes in forward scatter

Activation of neutrophils by fMLP results in a shape change that can be measured as change in forward scatter in a flow cytometer (26). Neutrophils (90 µl of a 2 x 10⁶ c/ml suspension) were incubated for 10 min at 37°C in a shaking water bath together with 10 µl of RPMI 1640-HSA or inhibitory protein (FLIPr or CHIPS). Subsequently, different concentrations of 10 times concentrated stimulus were added, and the cells were incubated for another 15 min at 37°C. The cells were finally fixed with an equal volume of 2.5% glutaraldehyde (Merck) in saline, and kept on ice for at least 90 min before measurement in a flow cytometer. After appropriate gating to exclude cell debris, the forward scatter values were determined.

Chemotaxis assays

Chemotaxis of human neutrophils toward several chemoattractants was measured in a 96-multiwell trans membrane system (ChemoTX; Neuro Probe) using an 8-µm pore size polycarbonate membrane. Cells (5 x 10⁶/ml) were labeled with 2 µM calcein-AM for 20 min at room temperature protected from light. Subsequently, cells were washed with HBSS containing 1% HSA, resuspended to 2.5 x 10⁶ cells/ml in the same buffer, and incubated with FLIPr. Dilutions of the different chemoattractants were prepared in HBSS-HSA, and 29 µl were placed into each well of the lower compartment of the chamber in triplicate. Wells with control medium were included to measure the spontaneous cell migration. For total cell fluorescence, wells were filled with 25 µl of labeled cells plus 4 µl of buffer. The membrane holder was assembled, and 25 µl of labeled cells were added as a droplet to each upper well except for the total fluorescence wells. The plate was incubated for 30 min at 37°C in a humidified 5% CO₂ atmosphere. The membrane was washed extensively with PBS and fluorescence of the wells was measured in a FlexStation fluorescent plate reader (Molecular Devices) with excitation at 485 nm and emission at 530 nm. The percentage of chemotaxis was calculated relative to the fluorescence value of cells added directly to the lower well: (fluorescence sample/fluorescence total counts) x 100.

Actin polymerization

To measure the polymerization state of actin in neutrophils after proper stimulation, a flow cytometric assay was performed using fluorescent phallacidin as probe, which binds specifically to F-actin, the active state of actin. A set of tubes was prepared with 25 µl of fixation/permeabilization buffer (6% formaldehyde in PBS with 200 µg/ml L--lysophosphatidylcholine). Neutrophils (5 x 10⁶ cells/ml) with or without inhibitor were stimulated at room temperature with LTB₄. The first sample (25 µl) was immediately added to a tube with fixation buffer, and consecutive samples at different time points. After keeping the samples for at least 15 min for fixation and permeabilization, 2 µl of the fluorescent probe (Alexa Fluo 488 phallacidin, 100 U/ml in methanol) was added. Samples were then kept at 4°C for 1 h and subsequently the fluorescence was measured on a flow cytometer.

Binding assay with leukocytes

To determine the binding of FLIPr to different leukocytes, isolated fractions of PMN and PBMC suspension were mixed again (4:6 ratio) and diluted to 5 x 10⁶ cells/ml with RPMI 1640 containing 1% HSA. Cells were incubated with buffer or a concentration range of FITC-labeled protein during 30 min. Cells were then washed and resuspended in RPMI 1640-HSA and binding of FLIPr was measured by flow cytometry. For binding in whole blood, 50 µl of EDTA anti-coagulated blood was incubated with 5 µl of different concentrations of FITC-labeled protein for 30 min at 4°C. Subsequently, samples were treated with FACS lysing solution, washed once, and the cells were resuspended in 200 µl of RPMI 1640-HSA and measured in the flow cytometer. The same protocol was also used for isolated PBMC adding the appropriate mAbs against different subsets of leukocytes, labeled with fluorochromes distinct from FITC: CD3-Cy5 plus CD4-PE or CD8-PE for T lymphocytes; CD19-PE for B lymphocytes; CD14-PE for monocytes; CD3-Cy5 plus CD56-PE and CD16-PE for NK cells.

Binding assay with HEK293

Cells transfected with each FLAG-tagged C5aR, FPR, and FPRL1 or 3xHA-tagged FPRL2 were incubated with mouse anti-FLAG or anti-HA mAb (10 µg/ml) for 45 min at 4°C. Cells were then washed and incubated with allophycocyanin-labeled goat anti-mouse Ab together with FITC-labeled FLIPr or CHIPS for 45 min at 4°C. Finally, the cells were washed and resuspended in 200 µl of RPMI-HSA containing 5 µg/ml propidium iodide. Association of FITC-protein (FL1) was determined to propidium iodide-negative living cells (scatters plus FL2) expressing the allophycocyanin-positive tagged receptor (FL4) in a flow cytometer (24). For background signals, cells transfected with an empty pcDNA3.1 vector were used.

	Results

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Prevalence in S. aureus isolates

To investigate the prevalence of the gene for FLIPr (designated flr) in clinical isolates, 91 S. aureus strains isolated from bloodstream infections were screened by PCR. The gene encoding for FLIPr was found in 59% of the isolates and encodes for an excreted protein that includes a signal peptide and AXA motive. The presence of FLIPr in the supernatants of growing S. aureus could not be directly determined due to the lack of specific Abs. Healthy blood bank donors possess natural occurring IgG Abs reacting with the rFLIPr indicating that FLIPr is produced in vivo (data not shown).

FLIPr inhibits FPR family-related activation of neutrophils

We first examined the capacity of FLIPr to inhibit cell responses to peptides reactive with members of the FPR family as measured by intracellular calcium mobilization. FLIPr itself, used as stimulus up to 100 µg/ml, did not induce a calcium response. Neutrophils were tested for activation with and without preincubation with 3 µg/ml FLIPr or CHIPS. Incubation of human neutrophils with FLIPr resulted in the inhibition of fMLP-induced calcium mobilization (Fig. 1A). Compared with CHIPS, the inhibition of fMLP-induced responses was weaker. The maximum inhibition of neutrophil activation by FLIPr was observed at a concentration of 3 x 10^–9 M fMLP, while CHIPS inhibited up to 10^–6 M fMLP. Because FLIPr partly inhibited the fMLP-induced activation of neutrophils, its activity was also tested on the low-affinity receptor FPRL1. Several synthetic peptides derived from a random peptide library, which have been reported as agonists of FPRL1 (10, 19, 20, 21, 27), were tested. A very strong inhibition of the FPRL1-specific MMK-1 peptide-induced activation of FLIPr-treated neutrophils was observed (Fig. 1C). FLIPr also inhibited WKYMVm- (FPR and FPRL1 agonist) and WKYMVM- (FPRL1 and FPRL2 agonist) induced responses in neutrophils (Fig. 1, B and D). The inhibition was stronger for WKYMVM. Although FLIPr inhibited the response to concentrations of 10^–8 M WKYMVm, it is able to inhibit up to 3 x 10^–7 M when using WKYMVM. CHIPS did not show any inhibitory activity in the response to FPRL1 agonists, as reported before (5). Unlike CHIPS, FLIPr did not block C5a-induced activation of neutrophils. In addition, FLIPr did not affect the response to other chemoattractant receptors present on neutrophils: LTB₄, PAF, IL-8, and GRO- (data not shown).

View larger version (20K): [in this window] [in a new window]   FIGURE 1. FLIPr inhibits FPR and FPRL1 agonist-induced calcium mobilization in neutrophils. Fluo-3-loaded neutrophils were incubated with buffer (•), 3 µg/ml FLIPr (), or CHIPS () for 20 min at room temperature. For calcium mobilization cells, each sample was first measured for 10 s to determine the basal fluorescence and subsequently stimulus was added and rapidly placed back in the sample holder to continue the measurement. Cells were analyzed in a flow cytometer and activation was expressed as the ratio of the fluorescence value before (cells acquired between T = 5 and 7 s)/after addition of stimulus (cells acquired at T = 12 until 14 s after stimulation). Neutrophils were stimulated with the synthetic agonists fMLP (A), WKYMVm (B), MMK-1 (C), and WKYMVM (D). Data are mean ± SEM of three independent experiments.

Monocytes also bear the receptors of the FPR family including the FPR, FPRL1, and FPRL2 that is not present on neutrophils (11, 13). The same set of agonists was used to stimulate the monocyte intracellular calcium mobilization in the presence of FLIPr or CHIPS. Specific monocyte response in the PBMC preparation was established by gating on side scatter and anti-CD14 staining. As shown in Fig. 2, FLIPr efficiently inhibited the response induced by MMK-1 (Fig. 2C, specific for FPRL-1), both WKYMVm (Fig. 2B) and WKYMVM (Fig. 2D). CHIPS did not affect these responses. Monocytes showed a smaller range of fMLP concentrations that induced activation as compared with the response induced in neutrophils (Fig. 1A). Only CHIPS and not FLIPr inhibited the fMLP-induced calcium mobilization in monocytes.

View larger version (18K): [in this window] [in a new window]   FIGURE 2. FLIPr inhibits FPR and FPRL1 agonist-induced calcium mobilization in monocytes. The activity of FLIPr was tested in calcium mobilization assays with monocytes in response to the synthetic agonists fMLP (A), WKYMVm (B), MMK-1 (C), and WKYMVM (D). Fluo-3-loaded PBMC were incubated with buffer (•), 3 µg/ml FLIPr (), or CHIPS () for 20 min and specific monocyte response determined by gating on anti-CD14-PE staining and scatter parameters. Data are mean ± SEM of three independent experiments.

To further investigate the potency of FLIPr, an experiment was performed with a dose response of both FLIPr and MMK-1. The effect was dose dependent and FLIPr inhibited the response to MMK-1 in the nanomolar to micromolar range (Fig. 3A). To further strengthen the specific inhibition of the fMLP-indued neutrophil activation, the change in forward scatter was monitored as an alternative parameter (26). CHIPS and FLIPr inhibited the fMLP-induced increase in forward scatter (Fig. 3B) in a manner similar to the inhibition of intracellular calcium mobilization (Fig. 1A). To support the effect of FLIPr on FPRL1 signaling, we examined its effect on neutrophil degranulation induced by MMK-1. Stimulation of cytochalasin-B-treated cells for 30 min with 10^–6 M MMK-1 resulted in the release of elastase (68% of total cell content) and myeloperoxidase (MPO; 11% of total cell content) as determined with chromogenic substrates (28). Pretreatment with 3 µg/ml FLIPr blocked the elastase release for 74% and the MPO release for 100%. CHIPS was unable to prevent the MMK-1-induced elastase and MPO release.

View larger version (10K): [in this window] [in a new window]   FIGURE 3. Potency of FLIPr to inhibit the MMK-1-induced calcium mobilization and fMLP-induced change in forward scatter of neutrophils. A, The activity of different concentrations FLIPr (0.01–10 µg/ml for 20 min) was tested in calcium mobilization assay with neutrophils in response to synthetic peptide FPRL1 agonist MMK-1 (concentration of 0 represents buffer-treated cells). A representative experiment is shown. B, Neutrophils were incubated with buffer (•), 3 µg/ml FLIPr (), or CHIPS () for 20 min at room temperature. Cells were challenged with different concentrations fMLP for 15 min at 37°C, fixed with 1% paraformaldehyde, and analyzed for the relative change in forward scatter value as compared with control cells incubated in buffer only. A representative experiment is shown.

To assess whether FLIPr could also inhibit the chemotactic response, neutrophil migration in response to the chemoattractants C5a, fMLP, and MMK-1 was determined in a multiwell chemotaxis assay. In accordance with the calcium mobilization assays, FLIPr did not show any effect on C5a. However, FLIPr partly inhibited the chemotactic response to fMLP and showed complete inhibitory activity toward MMK-1 (Fig. 4).

View larger version (9K): [in this window] [in a new window]   FIGURE 4. FLIPr inhibits chemotaxis of neutrophils to fMLP and MMK-1 and not to C5a. Chemotaxis of human neutrophils toward several chemoattractants was measured in a multiwell transmembrane system. Cells were loaded with calcein and incubated with buffer (•) or 3 µg/ml FLIPr (). Dilutions of the chemoattractants C5a (A), fMLP (B), and MMK-1 (C) were placed to each well in triplicate and, after assembling the membrane holder, labeled cells were added to each upper well. The plate was incubated for 30 min at 37°C plus 5% CO2, and after washing the membrane holder, fluorescence was measured. Results are expressed as percentage of chemotaxis, and data are mean ± SEM of triplicates from one representative experiment of three. Spontaneous migration toward buffer loaded wells was 29%.

FLIPr inhibits A_1–42- and PrP_106–126-induced activation of neutrophils

The activation of monocyte-derived cells is thought to play a key role in the inflammatory process leading to the pathogenesis of many neurodegenerative diseases (29). Although the potential involvement of other cell surface receptors should not be excluded, FPRL1 has been proposed to mediate the migration and activation of monocytes and microglia induced both by A_1–42 (16) and by a 20-aa fragment of the human prion protein PrP_106–126 (17). Neutrophils are also activated by both A_1–42 (30, 31) and PrP_106–126 (32) leading to changes in intracellular calcium, generation of superoxide, and induction of chemotaxis. Therefore, we examined the capacity of FLIPr to inhibit the calcium mobilization in response to 10 µM A_1–42 and 50 µM PrP_106–126 (Fig. 5A). For comparison, the potent inhibition of MMK-1- and fMLP-induced calcium mobilization by FLIPr was performed in parallel. With A_1–42, a specific migration was induced that was partly inhibited by FLIPr (Fig. 5B). Because the A_1–42-induced calcium response as determined by Fluo-3 and flow cytometry were relatively weak, the experiment was repeated with fura-2-labeled cells and ratiometry in a fluorescent plate reader (FlexStation). This enabled a more clear view on the A_1–42-induced calcium response that was completely inhibited by FLIPr (Fig. 5C). To demonstrate specificity of the response, the same cells were rechallenged after 5 min with PAF. This elicited a calcium mobilization in all cells, both treated with buffer and FLIPr.

View larger version (10K): [in this window] [in a new window]   FIGURE 5. FLIPr inhibits chemotaxis and calcium flux in response to the endogenous peptide agonist A1–42 and PrP106–126. The activity of FLIPr to inhibit the neutrophil response to FPRL1-endogenous agonists A1–42 and PrP106–126 was tested by chemotaxis and calcium mobilization. A, The calcium flux induced by 10 µM A1–42 (AB) and 50 µM PrP106–126 (PrP) was inhibited by 3 µg/ml FLIPr. In the same experiment, the peptide agonists MMK-1 (1 x 10–7 M) and fMLP (1 x 10–9 M) were included. , The response of buffer control cells; , the response in the presence of FLIPr. B, Chemotaxis results toward different concentrations A1–42 of control cells (•) and cells incubated with 3 µg/ml FLIPr (). Data are expressed as percentage of migration and are mean ± SEM of triplicates of one representative experiment. Controls are included of chemotaxis in response to 3 x 10–7 M MMK-1 in control cells () and in cells incubated with FLIPr (). Spontaneous migration toward buffer was 21.8%. C, Representative experiments showing A1–42 (10–5 M at 60 s) induced calcium mobilization in fura-2-loaded neutrophils treated with buffer, or 3 µg/ml FLIPr. The same cells were rechallenged at 300 s with 10–9 M PAF. Results are depicted as the ratio of the fluorescence at 530/590 nm and shifted to show the individual curves.

Lipoxin A4 is an endogenous lipid-derived mediator generated at sites of inflammation that has been reported to bind FPRL1/LXA4R with high affinity (33). Unlike peptide chemotactic agonists, lipoxin A4 induces an anti-inflammatory signaling cascade that inhibits neutrophils migration (34) and suppresses calcium mobilization upon challenge with other agonists (35). Lipoxin A4 was also tested as a direct FPRL1 agonist in the calcium mobilization assay. However, we were unable to elicit a calcium response in neutrophils or monocytes in response to fresh lipoxin A4; neither when assayed with Fluo-3 and flow cytometry nor with fura-2 and ratiometry in a fluorescent plate reader. To investigate a possible antagonistic effect of FLIPr for lipoxin A4, inhibition of LTB₄-induced actin polymerization was measured. Cells incubated with 10^–6 M lipoxin A4 showed a decreased actin polymerization in response to LTB₄. Preincubation with FLIPr at different concentrations could not revert this effect. FLIPr itself did not inhibit the actin polymerization in response to LTB₄, in accordance with the results obtained with calcium mobilization (Fig. 6).

View larger version (16K): [in this window] [in a new window]   FIGURE 6. FLIPr does not interfere with lipoxin A4-mediated FPRL1 activation. The LTB4-induced (10–9 M) actin polymerization is partly prevented by the incubation of neutrophils with 10–6 M lipoxin A4. Preincubation of neutrophils with 3 µg/ml FLIPr did not interfere with the LTB4-induced response nor the lipoxin A4 response. Actin polymerization was determined at 15 s intervals with Alexa-labeled phallacidin and flow cytometry for cells plus LTB4 (•), FLIPr and LTB4 (), lipoxin (LPX) A4 and LTB4 (), and FLIPr plus lipoxin A4 and LTB4 (dashed line, ). Results are expressed as the relative increase in fluorescence compared with nonstimulated cells (mean of two representative experiments).

To show association of FLIPr with the appropriate blood leukocytes that bear FPRL1, fluorescent-labeled FLIPr was used. With neutrophils and monocytes, a strong association of FLIPr-FITC was observed, while lymphocytes showed a weak binding (Fig. 7). With increasing concentrations of FLIPr-FITC, an increase in binding was observed, both when cells were incubated at 37°C (Fig. 7A) and on ice (Fig. 7B). To test whether binding was influenced by plasma component, the experiment was also performed using whole blood ex vivo. The results were not different from binding to isolated leukocytes (data not shown). mAbs against different PBMC subtypes were used together with FLIPr-FITC to determine the binding profile of FLIPr to different cell populations (Fig. 8). Binding was observed to monocytes (CD14⁺, gated on scatters), B cells (CD19⁺ lymphocytes), a subpopulation of CD8⁺ lymphocytes and NK cells (CD3^–/CD56⁺/CD16⁺ lymphocytes). The CD8⁺ subpopulation that bound FLIPr was identified as NK cells (CD56⁺,CD8⁺). No binding was found to T cells (CD3⁺ lymphocytes), or the CD4⁺ subset and the majority of CD8⁺ subset. The FITC-labeled FLIPr was also functional in a calcium mobilization assay (using fura red instead of Fluo-3-AM) inhibiting fMLP-, WKYMVm- and MMK-1-induced activation of neutrophils (data not shown).

View larger version (9K): [in this window] [in a new window]   FIGURE 7. FLIPr binds to neutrophils, monocytes, and a proportion of lymphocytes. Isolated PMN and PBMC were incubated with a range of concentrations of FLIPr-FITC (0.03–9 µg/ml) for 30 min on ice (A) or at 37°C (B) under constant shaking. Cells were then washed and resuspended in RPMI 1640-HSA and fluorescence was measured in a flow cytometer. Cells were identified based on scatter parameters and anti-CD14 staining; neutrophils (•), monocytes (), and lymphocytes () are displayed. Data are mean ± SEM of three independent experiments.

View larger version (20K): [in this window] [in a new window]   FIGURE 8. FLIPr binds to different subsets of leukocytes. mAbs for different subsets of mononuclear cells were used to check the binding profile of FLIPr-FITC by flow cytometry. FLIPr binds to: CD14+ monocytes (A), not to CD3+ lymphocytes (T cells) (B); binds to CD19+ lymphocytes (B cells) (C), not to CD4+ T cells (D); binds to a subpopulation of CD8+ T cells (E), and to CD3–CD56+CD16+ lymphocytes (NK cells) (F).

To assess whether FLIPr binds directly to the human receptor FPR and/or FPRL1, HEK293 cells transiently transfected with FLAG-tagged FPR and FPRL1 were tested for FLIPr-FITC binding. As positive controls, CHIPS-FITC binding and C5aR-transfected HEK293 were included. Cells were analyzed by gating on forward and side scatters as well as viability (cells staining negative for propidium iodide) to exclude dead cells. Indirect allophycocyanin-labeled mAb against the FLAG or 3xHA tag detected the population of transfectants expressing the respective receptors. Fig. 9A shows representative histograms of the binding of FLIPr-FITC and CHIPS-FITC to the transfectants. As expected, CHIPS-FITC (3 µg/ml) bound to HEK293 transfected with FPR as well as those transfected with C5aR and did not bind to cells transfected with FPRL1. FLIPr-FITC (3 µg/ml) bound to HEK293 transfected with FPRL1, did not bind to HEK293 transfected with C5aR or FPRL2 and showed a weak binding to cells transfected with FPR. Binding to vector-control transfectants gave a mean fluorescence of 8.6 ± 1.1 (Fig. 9B).

View larger version (21K): [in this window] [in a new window]   FIGURE 9. FLIPr binds to HEK293 cells transfected with the FPRL1. HEK293 cells were transiently transfected with the vector containing FLAG-tagged human FPR, FPRL1, and C5aR or 3xHA-tagged FPRL2. As control, an empty vector was used. To identify positive transfectants, cells were labeled with anti-FLAG mAb (or anti-HA mAb for FPRL2) and allophycocyanin-labeled goat anti-mouse IgG Ab. Simultaneously, FITC-labeled FLIPr or CHIPS was added at 3 µg/ml. Cells were resuspended in buffer with propidium iodide and analyzed for binding of FITC-labeled protein to viable, receptor-positive transfectants. Therefore, cells were gated on basis of scatters and viability (propidium iodide negative) and analyzed for expression of the receptor on the cell surface (allophycocyanin positive) and binding of FITC-labeled protein. A, Representative histograms of the binding of CHIPS-FITC to C5aR, FPR, and FPRL1 (left column) and FLIPr-FITC to C5aR, FPR, FPRL1, and FPRL2 (right column). Background staining to vector control cells is depicted as gray overlays. B, The mean fluorescence ± SEM of three independent experiments; , FLIPr-FITC; , CHIPS-FITC binding. Mean fluorescence value for binding to vector control HEK293 cells was 8.6 ± 1.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

The role of FLIPr, as an FPRL1 inhibitor, in the pathogenesis of staphylococcal disease is difficult to assess, but several data make the protein likely to play a role in host-pathogen interactions. The gene encoding for FLIPr was found to be located in a genetic cluster which contains genes encoding several virulence factors: extracellular fibrinogen-binding protein (efb), extracellular fibrinogen-binding protein-like (efb-L), hemotoxin protein A (better known as -toxine, hla), and enterotoxin-like proteins as well as an insertional sequence (tnp IS1181). Furthermore, the gene is present in 59% of clinical isolates. The gene for FLIPr encodes for a protein containing a leader and an AXA motive as previously described for CHIPS (5).

The role of C5a and fMLP in host defense is clear, because they have been implicated not only in leukocyte recruitment but also on the hepatic synthesis of acute phase proteins (8), as well as activation of NF-B and production of inflammatory cytokines by phagocytes (37). Furthermore, targeted gene disruption of FPR in mice impaired the antibacterial response (38).

Less is known about FPRL1, but there is increasing evidence of a critical role in the regulation of immune responses against infections. Not only fMLP but several microbial components as well as host molecules implicated in innate defense are able to activate FPRL1. Activation by several agonists has shown its ability to mediate chemotaxis as well as superoxide generation by phagocytes (13). Activation of FPRL1 by domains of the HIV envelope proteins has been reported to desensitize chemokine receptors CCR5 and CXCR4 that act as coreceptors for HIV infection (39, 40). Also a cecropin-like Helicobacter pylori peptide, Hp_2–20, attracts and activates monocytes through FPRL1 and FPRL2 (13, 41).

In contrast, LL-37, an enzymatic cleavage fragment of the neutrophil granule-derived cathelicidin, an endogenous antimicrobial peptide, uses FPRL1 as a receptor to activate human neutrophils, monocytes, and T cells (42). In addition, LL-37 also prolongs the lifespan of neutrophils by suppression of apoptosis mediated via activation of FPRL1 (43). LL-37 is an important effector molecule of the innate immune system which possesses considerable antistaphylococcal activity and may contribute to protection of the skin and mucosal surfaces against colonization by Gram-positive pathogens (44). The production of aureolysin, a metalloproteinase that cleaves and inactivates LL-37, by S. aureus contributes to the resistance against bactericidal peptides (45). Several other mechanisms are used in protection against cationic antimicrobial molecules such as defensins (46). A truncated form of a -chemokine has also been reported as a high-affinity FPRL1 agonist (47). FPRL1 also interacts with the lipid metabolite lipoxin A4, which plays an anti-inflammatory role. Lipoxin A4 has been reported to induce calcium mobilization in neutrophils and monocytes in some studies (48, 49), while in others it could not induce any direct cellular response (21, 47). In our experience, lipoxin A4 did not elicit a calcium mobilization but did inhibit actin polymerization in response to LTB₄, which could not be reverted by FLIPr. It is hypothesized that lipoxin A4 and peptide agonists might use divergent domains on FPRL1 and differential cellular signaling (48). FPRL1 recognizes a vast array of ligands that have no homology; different functional domains in the receptor are probably used for diverse agonists, as it has been demonstrated for MMK-1 and A_1–42 (50). Thus, FLIPr is not able to block the response to every FPRL1 agonist.

A better understanding of the virulence strategies of S. aureus might lead to development of more specific therapeutically approaches. Some virulence factors have been associated with specific clinical features, such as the staphylococcal syndromes associated with toxin production (1). In contrast, while some factors are present in strains causing disease by direct invasion and tissue destruction, others are associated with secondary invasive infections as infective endocarditis. Whether an infection is contained or spreads depends on a complex interplay between S. aureus virulence determinants and host defense mechanisms (1, 2). The blocking of receptors for chemoattractants exerted by the staphylococcal proteins CHIPS and FLIPr may have a role in preventing the early detection of the microorganism by the innate immune mechanisms, allowing its spread.

Leukocyte migration is critical in maintaining the host defense, aiming at the clearance of noxious agents. Uncontrolled cellular infiltration into tissues can lead to chronic inflammation and toxic release of substances such as superoxide anions. FPRL1 may constitute an important molecular target for the development of new therapeutic agents to combat excessive inflammatory responses. The activation of FPRL1 by A_1–42 or PrP_106–126 may be responsible for accumulation and activation of mononuclear phagocytes (monocytes and microglia) as well as fibrillar formation that is associated with the pathogenesis of Alzheimer’s disease and prion diseases, respectively (18). FPRL1 has been shown to participate in the process of endocytosis and subsequent aggregation of A_1–42 in mononuclear phagocytes (51). It has also been described that the endogenous peptide humanin would exert its neuroprotective effects by competitively inhibiting the access of A_1–42 to FPRL1 (52). Nevertheless, not only FPRL1 but also a number of other putative surface receptors have been described to interact with A_1–42 (53). The Alzheimer’s patient will benefit from a combination of different drugs and the development of FPRL1-specific antagonists may have promising therapeutic potential in retarding the progression of the disease.

The leukocyte-binding profile of FLIPr is also intriguing. In our experiments, FLIPr-FITC binding was observed to monocytes, neutrophils, B cells, a subpopulation of CD8⁺ lymphocytes (most likely the NKT cells), and also very clearly to NK cells. Monocytes express more FPRL1 than neutrophils, but less is reported for lymphocytes. Human CD3⁺ T lymphocytes showed a weak migration in response to high concentrations of T21/DP107, which could be mediated by the presence of FPRL1 (54). NK cells express receptors for several chemoattractants. They migrate toward concentration gradients of C5a and fMLP, and chemokines also appear to play an important role in afferent and efferent NK cell responses to both infected and neoplastic cells (55). The expression of receptors for CXC chemokines, CC chemokines, and CX3C chemokines has been described (56). Presence of FPRL1 on NK cells has not been described, although the effect of the FPRL1 agonist lipoxin A4 on NK cells has been reported previously (57). In contrast, just as CHIPS was found to bind and functionally inhibit two different G protein-coupled receptors (5), it is not unlikely that FLIPr also binds to a receptor other than FPRL1.

	Disclosures

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The authors have no financial conflict of interest.

	Footnotes

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

¹ This work was supported by grants "Fondo de Investigacion Sanitaria, Instituto de Salud Carlos III (expediente 01/F062)" and "Sociedad Española de Enfermedades Infecciosas y Microbiologia Clinica," (both to C.P.), and by the European Union (LSHM-CT-2004-512093) and the Netherlands Genomics Initiative (050-71-249).

² Address correspondence and reprint requests to Dr. Kok P. M. van Kessel, Eijkman-Winkler Institute, University Medical Center Utrecht, G04.614, Heidelberglaan 100, 3584 CX Utrecht, The Netherlands. E-mail address: k.kessel{at}umcutrecht.nl

³ Abbreviations used in this paper: LTB₄, leukotriene B₄; PAF, platelet-activating factor; CHIPS, chemotaxis inhibitory protein of S. aureus; FPR, formyl peptide receptor; FPRL, FPR-like receptor; A, amyloid ; FLIPr, FPRL1 inhibitory protein; HA, hemagglutinin; PMN, polymorphonuclear neutrophil; HSA, human serum albumin; HEK, human embryonic kidney; GRO, growth-related oncogene; MPO, myeloperoxidase.

Received for publication March 24, 2006. Accepted for publication August 30, 2006.

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