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The Journal of Immunology, 2007, 179, 2520 -2531
Copyright © 2007 by The American Association of Immunologists, Inc.
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C-Terminal Tail Phosphorylation of N-Formyl Peptide Receptor: Differential Recognition of Two Neutrophil Chemoattractant Receptors by Monoclonal Antibodies NFPR1 and NFPR21

Marcia Riesselman*, Heini M. Miettinen*, Jeannie M. Gripentrog*, Connie I. Lord*, Brendan Mumey{ddagger}, Edward A. Dratz{dagger}, Jamal Stie*, Ross M. Taylor* and Algirdas J. Jesaitis2,{dagger}

* Departments of Microbiology, Chemistry and {dagger} Biochemisty, and {ddagger} Computer Science, Montana State University, Bozeman, Montana 59717


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The N-formyl peptide receptor (FPR), a G protein-coupled receptor that binds proinflammatory chemoattractant peptides, serves as a model receptor for leukocyte chemotaxis. Recombinant histidine-tagged FPR (rHis-FPR) was purified in lysophosphatidyl glycerol (LPG) by Ni2+-NTA agarose chromatography to >95% purity with high yield. MALDI-TOF mass analysis (>36% sequence coverage) and immunoblotting confirmed the identity as FPR. The rHis-FPR served as an immunogen for the production of 2 mAbs, NFPR1 and NFPR2, that epitope map to the FPR C-terminal tail sequences, 305-GQDFRERLI-313 and 337-NSTLPSAEVE-346, respectively. Both mAbs specifically immunoblotted rHis-FPR and recombinant FPR (rFPR) expressed in Chinese hamster ovary cells. NFPR1 also recognized recombinant FPRL1, specifically expressed in mouse L fibroblasts. In human neutrophil membranes, both Abs labeled a 45–75 kDa species (peak Mr ~60 kDa) localized primarily in the plasma membrane with a minor component in the lactoferrin-enriched intracellular fractions, consistent with FPR size and localization. NFPR1 also recognized a band of Mr ~40 kDa localized, in equal proportions to the plasma membrane and lactoferrin-enriched fractions, consistent with FPRL1 size and localization. Only NFPR2 was capable of immunoprecipitation of rFPR in detergent extracts. The recognition of rFPR by NFPR2 is lost after exposure of cellular rFPR to f-Met-Leu-Phe (fMLF) and regained after alkaline phosphatase treatment of rFPR-bearing membranes. In neutrophils, NFPR2 immunofluorescence was lost upon fMLF stimulation. Immunoblotting ~60 kDa species, after phosphatase treatment of fMLF-stimulated neutrophil membranes, was also enhanced. We conclude that the region 337–346 of FPR becomes phosphorylated after fMLF activation of rFPR-expressing Chinese hamster ovary cells and neutrophils.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The N-formyl peptide receptor (FPR)3 is a G protein-coupled receptor (GPCR), found in leukocytes, that mediates host defensive responses including chemotaxis, secretion, and superoxide production (1, 2, 3). The human receptor is in low abundance and has yet to be purified (4) in quantities suitable for biochemical or structural characterization. As most GPCRs thus far examined, FPR activity is exquisitely regulated by cellular kinases (5, 6, 7, 8), surface reorganization (9, 10), internalization and recycling (11) and interaction with adaptor proteins such as arrestins (12) and possibly others (13). These forms of regulation may rely on receptor posttranslational modifications, conformational changes, and/or spatial accessibility to signal transduction partners.

It is now well known that, most if not all, GPCRs have cytoplasmic loci of phosphorylation (14, 15) that apparently determine cellular fates before and after occupancy of ligand. To date, most of the studies that have examined phosphorylation sites on FPR and the consequences of their modulation have been performed in model cell culture systems (5, 16, 17). Such systems may or may not serve as adequate models of the neutrophil and macrophage, where this receptor is normally found. Furthermore, besides structural analysis of rhodopsin and bacteriorhodopsin, x-ray crystallography has not been successfully applied for other GPCRs, nor has analysis of the activated structure of any GPCR been elucidated. Additionally, monoclonal immunoprobes of denatured or ligand occupied FPR are not available, significantly impeding the biochemical analysis of the occupied state.

To facilitate further cell biological and biochemical analysis of FPR, we have expressed and purified a hexa-histidine-tagged recombinant FPR (rHis-FPR) in Spodoptera frugiperda (Sf9) cells and used it as an immunogen to generate mAbs to FPR. The identity of purified material was confirmed by mass analysis of tryptic peptides and by immunoblot analysis, using monoclonal anti-His-tag or rabbit anti-FPR peptide polyclonal Abs. The epitopes of the two mAbs generated from the rHis-FPR immunogen were mapped by phage display technology, suggesting that these Abs recognize specific regions of the cytoplasmic face of FPR. This identification accounted for their high sensitivity in recognizing rHis-FPR, rFPR expressed in Chinese hamster ovary (CHO) cells, and a molecular species consistent with human neutrophil FPR. The NFPR2 Ab was able to immunoprecipitate the FPR in dodecyl maltoside membrane extracts and was sensitive to activation state of the cells from which FPR was obtained. The NFPR1 Ab also recognized the closely related chemoattractant receptor FPRL1. Both Abs produced immunofluorescence images consistent with receptor enrichment in the plasma membrane- and intracellular-specific/gelatinase granule-enriched fractions of neutrophils.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Materials

The pBlueBacHisB transfer vector was obtained from Invitrogen Life Technologies. Dodecylmaltoside was obtained from Anatrace; pentadecafluorooctanoic acid was purchased from Fluka; octylglucoside, PMSF, and DTT from Calbiochem; and lysophosphatidyl glycerol (LPG) from Avanti. The anti-His-tag mAb was purchased from Novagen. Lactoferrin polyclonal Ab was obtained from ICN Pharmaceuticals. Goat anti-mouse (H+L)-conjugated with Alexa Fluor 488 was purchased from Molecular Probes. CNBr-activated Sepharose 4B, protein A-Sepharose and GammaBind Plus-Sepharose beads were obtained from Pharmacia; Ni2+-NTA agarose from Qiagen; BioMax and Centricon concentrators from Millipore; and the goat-anti-rabbit (H+L) alkaline phosphatase, goat-anti-mouse (H+L) alkaline phosphatase, FITC secondary Ab, and Econo-Pac 10 DG desalting columns (30 x 10 ml) from BioRad. ProSieve (Cambrex) color m.w. markers were purchased from BMA; isopropyl beta-D-thiogalactoside from Promega; immunoblot developing reagents (NBT/BCIP) were purchased from Kirkegaard & Perry Laboratories; and nitrocellulose (0.45 µm pore size) from Schleicher & Schuell. Powder for Luria-Bertani broth (LB) and LB agar was obtained from VWR International. SDS, 0.2 µm polyethersulfone syringe filters (Whatman), acrylamide, bis-acrylamide, NaCl, NaH2PO4, KCl, EDTA, EGTA, NaN3, chloroform, and HPLC grade methanol were purchased from Fisher Scientific. The BCA protein determination and Supersignal chemiluminescence detection kits were obtained from Pierce Biotechnology and Maxisorb 96-well ELISA plates from Corning. The anti-rhodopsin mAb K16 was produced in-house by standard hybridoma culture technology from hybridoma cells provided by Prof. Paul Hargrave (University of Florida College of Medicine, Gainesville, FL). TX6 and TX31 mouse L cell fibroblast lines were provided by Prof. Richard Ye (University of Illinois College of Medicine, Chicago, IL). Trypsin Gold and shrimp alkaline phosphatase was obtained from Promega; and acetonitrile, isopropanol, and trifluoroacetic acid from J.T. Baker. The MALDI matrix {alpha}-cyano-4-hydroxycinnamic acid was purchased from Sigma-Aldrich; 5-bromo-4-chloro-3-indolyl-beta-D-galactopyranoside Bluo-Gal from Invitrogen Life Technologies; and Immobilon-P polyvinylidene fluoride from Millipore and Bio-Rad. Fix & Perm was purchased from Caltag Laboratories. Polyethylene glycol was obtained from Roche Pharmaceuticals. All other reagents were obtained from Sigma-Aldrich.

Cellular FPR in Sf9, CHO, and human neutrophils

FPR cDNA (18) was amplified by PCR with primers containing BamHI and HindIII restriction sites. Following digestion and purification, the PCR product was ligated into the BamHI-HindIII cloning sites of pBlueBacHis B and the sequence was confirmed by dideoxy-sequencing. The vector was transfected into Sf9 insect cells with linear AcMNPV DNA using the cationic liposome method (19). A virus stock, collected 48 h posttransfection, was used for infection of Sf9 cells and plaque purification of recombinant virus according to the manufacturer’s instructions. The expression of rHis-FPR in Sf9 cells infected with recombinant virus was confirmed by immunofluorescence (not shown) and immunoblots, after which high titer virus stocks were prepared. CHO cells, expressing recombinant FPR without a His tag (20), were also used as a source of rFPR. Human neutrophils were prepared from whole blood as previously described (21) and served as the source of native FPR.

Preparation of membranes and detergent extracts of rHis-FPR, rFPR, FPR, and rFPRL1

Suspension cultures of Sf9 cells were grown to 1–2 x 106 cells per millilter at 28°C in TNM-FH medium supplemented with 10% FBS, 50 µg/ml gentamicin, and 0.1% pluronic F-68 as described (22). Pelleted cells (1000 x g for 10 min) were suspended in fresh medium and inoculated at ~2 x 107/ml with rHis-FPR baculovirus stock at a multiplicity of infection of 5 for 1 h at 28°C. The cell suspension was returned to 1 L spinner flasks at 1–2 x 106/ml with fresh medium and harvested 60 h post-infection by washing in sterile saline at 1000 x g. The cell pellet was weighed and resuspended in 10 vol of N2 cavitation buffer (10 mM HEPES, pH 7.4, 100 mM KCl, 10 mM NaCl, 1 mM DTT, 3.5 M MgCl2, 1 mM ATP, 0.5 mM PMSF, 10 µg/ml aprotinin, 1 mM benzamidine, and 1 µl of protease inhibitor mixture per milliliter (Sigma P8340), and the cells disrupted as previously described for neutrophils (23). The crude cavitates (with 1 mM EGTA added) were centrifuged at 1000 x g for 10 min to produce the low speed supernatant and pellets 1KS and 1KP, respectively. The 1KS fraction was then centrifuged at 100,000 x g for 60 min and the pellets (100KP1) and supernatant (100KS1) saved for analysis. The 100KP1 was then resuspended in one half of the 1KS vol of 10 mM NaOH and incubated on ice for 10 min, to strip membrane skeletal and peripheral membrane proteins from the membranes as previously described for neutrophils (24). These extracts were then centrifuged again at 100,000 x g for 30 min to generate the second set of 100K pellets and supernatants, 100KP2 and 100KS2. Membranes of wild-type and FPR-expressing CHO cells, and human neutrophils were prepared as described by Miettinen et al. (20) and Parkos et al. (23), respectively and stored at –70°C. TX6 cells, expressing recombinant FPRL1 (rFPRL1) and transfection control (empty vector), TX31 cells (25), were released from culture dishes with Dulbecco’s PBS (DPBS), 1 mM EDTA, and membranes prepared as described for SF9 cells. Membranes were alkali stripped as described above.

Purification of rHis-FPR from Ni2+ affinity matrix

Membrane solubilization with LPG was carried out according to a modification of Huang et al. (26). Briefly, the 100KP2 was suspended in 25 mM HEPES (pH 8.0), 10% glycerol, and used fresh or stored at –70°C. Alkali-stripped membranes at 2 mg/ml protein were added to glass tubes with films of argon-dried LPG to achieve 10 mg/ml LPG with 1 mM 2-ME then stirred to extract for 1 h on ice. The extract was then centrifuged at 100,000 x g for 60 min to make the final set of pellet and supernatant fractions, 100KP3 and 100KS3. The 100KS3 is the soluble rHis-FPR extract. rHis-FPR containing LPG extract from 20 mg of membrane protein was tumbled overnight at 4°C with 2 ml of Ni2+-NTA agarose, prewashed two times with 6 ml of LPG column buffer (25 mM HEPES (pH 8.0), 10% glycerol, 400 mM NaCl, and 0.01% LPG). In a fritted disposable column, the loaded matrix was washed twice with 8 ml of column buffer containing 20 mM imidazole and suspended in 2 ml of this buffer and sampled for analysis by SDS-PAGE. rHis-FPR was eluted with six 1-ml volumes of LPG column buffer containing 300 mM imidazole. The final resin was suspended in 2 ml of buffer and again sampled for analysis by SDS-PAGE.

SDS-PAGE and immunoblot analysis

Membrane and purification samples were mixed with an equal volume of SDS-PAGE sample buffer (3.3% (w/v) SDS, 167 mM Tris-HCl (pH 6.8), 33% (v/v) glycerol, 0.03% (w/v) bromphenol blue, and 500 mM 2-ME). The samples were separated by SDS-PAGE on 9% or 7–15% Tris/glycine polyacrylamide slab gels containing 0.1% (w/v) SDS as described (27) and electrophoretic mobility of the sample proteins was calibrated using Prosieve prestained standards. Proteins were visualized on slab gels by staining with 0.125% Coomassie blue G 250 in 50% methanol/10% acetic acid as described (28), silver stained under basic conditions as described (29) or were prepared for immunoblotting as described below. Following SDS-PAGE, proteins were electrotransferred onto Immobilon-P polyvinylidene fluoride or nitrocellulose membranes and prepared for immunoblot analysis. Membranes were first incubated in blocking solution of 5% nonfat dried milk in TBST (0.14 M NaCl, 20 mM Tris-HCl (pH 7.4), 0.05% Tween 20) for 1–16 h at ambient temperature or 4°C, followed by a brief wash in TBST, and then exposed to ~0.2 µg/ml Ab specific to the His-tag, affinity purified rabbit polyclonal Ab specific to the carboxyl-terminal tail peptide of FPR, anti-c-terminal 337-FPR-350 peptide (anti-CTZ) (1:200) (30), lactoferrin (1:1000), or mAbs (1:1000 or ~2–4 µg/ml) in blocking buffer or with 5% BSA replacing the milk. After 3 x 10 min washes in TBST, immunoblots were incubated with 1:1000 secondary alkaline phosphatase-conjugated goat anti-rabbit or goat anti-mouse Ab for 2–4 h at room temperature. Subsequently, the immunoblots were washed 3 more times and then developed using an alkaline phosphatase NBT/BCIP development kit. Alternatively, blots were also developed using peroxidase and chemiluminescence with photographic film to visualize FPR using Supersignal chemiluminescence detection kit. Colorimetric signal or densitometric signal was quantitated from optical scans of dried blots or chemiluminescence-exposed film using ONE-Dscan/1-D Gel Analysis software for windows (Scanalytics). The primary and secondary Ab concentrations used above were selected to yield OD readings in the linear range of detection for each Ab.

For certain experiments, whole CHO cell extracts were prepared by incubating cells on 35-mm dishes with different concentrations of fMLF or for different times. Cells were rinsed with cold PBS and 200 µl of hot (60°C) Laemmli sample buffer was added, and the cells were scraped off the dish. The cell extract was incubated for 20 min at 60°C and 8 µl (~3.2 x 104 cells) were then loaded per lane for SDS-PAGE analysis. The primary Abs were diluted at 3 µg/ml for the immunoblot.

Alkaline phosphatase treatment of membrane FPR

Alkali-stripped CHO or neutrophil membranes were prepared as described above and incubated with shrimp alkaline phosphatase to dephosphorylate integral membrane proteins, including FPR (31). 15 µg of membranes in 50 µl of dephosphorylation buffer were incubated with 10 U of shrimp alkaline phosphatase at 37°C for varying periods of time up to overnight (o/n, 16 h). At termination, reactions were diluted to 1 ml in iced buffer and the membranes pelleted, at which point 0.2 (chemiluminescence) or 0.5 µg (colorimetric) of membranes were solubilized in SDS-PAGE sample buffer, electrophoresed, and electrotransferred to nitrocellulose. These samples were then immunoblotted with NFPR1 and NFPR2 and developed as described above.

Production of monoclonal anti-FPR antibodies

Using standard hybridoma technology, mAbs NFPR1 and NFPR2 were prepared against rHisFPR. Ni2+-NTA agarose affinity purified rHis-FPR was diluted into 10 mM HEPES (pH 8.0), 100 mM KCl, 50 mM NaCl, pH 7.4, 0.01% LPG, and concentrated to 2 mg of protein/ml in a BioMax 10K NMWL concentrator. Residual glycerol was ~0.2–0.3% and imidazole ~4–5 mM in the final immunogen preparation. This immunogen was sterile filtered and ready for direct i.v. injection, or emulsified with IFA for i.p. administration. Female BALBc mice were immunized with initial i.p. doses of 100 µg of rHis-FPR quantitated colorimetrically using the Pierce BCA kit, and then boosted on days 14, 28, and 42 with 25 µg i.p. and 25 µg i.v. A mouse with high serum Ab titer against FPR on SDS-PAGE immunoblots of FPR expressing CHO cell membranes was given final immunization injections, and the spleen harvested for fusion 4 days later. Splenocytes were fused with P3U1 myeloma cells using 1500 MW polyethylene glycol, plated into hypoxanthine aminopterin thymidine (HAT) selection medium, and cultured as previously described (32). Hybridomas from 7 to 14 day culture supernatants that screened positive by ELISA for mouse IgG and against intact and permeabilized CHO-FPR monolayers (but negative against unpermeabilized monolayers and nonexpressing CHO cells) were confirmed on SDS-PAGE immunoblots of FPR expressing CHO cell membranes, and cloned twice by limiting dilution. These positive clones were expanded and mAb produced in culture supernatants was isolated as previously described (33).

Flow cytometry

Diisopropylfluorophosphate-treated human neutrophils, purified from blood as previously described (21, 34), were incubated for 5 min at 37°C with or without 1 µM fMLF at 107 cells per milliliter of DPBS, 0.1% BSA, and 0.1% dextrose. For each sample, 106 cells were fixed using Fix & Perm by the manufacturer’s methanol modification protocol, washed once with 5% BSA in DPBS, 0.1% azide, and suspended in 100 µl permeabilization medium. Primary or control Ab was added to 10 µg/ml and incubated for 30 min at ambient temperature. The cells were washed twice and incubated for 30 min in Alexa Fluor 488-conjugated goat anti-mouse secondary, diluted 1:4000 in wash buffer. Again, the cells were washed twice, suspended in 0.5 ml of DPBS and fluorescence intensity was determined on a Becton Dickinson FACScan analyzer.

Immunofluorescence microscopy

Diisopropylfluorophosphate-treated human neutrophils purified from blood by the gelatin method as above were incubated for 5 min at 37°C with or without 1 µM fMLF at 107 cells per milliliter of DPBS, 0.1% BSA, 0.1% dextrose, then diluted to 5 x 105 cells per milliliter. 100 µl of cell suspension was centrifuged to a glass slide at 200 rpm for 2 min in a Cytospin 2 (Shandon Southern Products). The cells were immediately fixed in methanol for 5 min, rinsed in two changes of DPBS and blocked in DPBS, 0.2% gelatin, 3% normal goat serum, 0.02% azide overnight at 4°C. For immunofluorescence, the cells were incubated for 1 h ambient temperature in 10 µg/ml mAb, washed in four changes of DPBS, 0.2% gelatin, 0.02% azide and once in blocking buffer with goat serum, then placed in a 1:4000 dilution of Alexa Fluor 488 conjugated goat anti-mouse secondary Ab and the incubation and wash series repeated; 90% glycerol was used to mount cover slips. The immunofluorescence-stained cells were evaluated and photographed with a Zeiss Axioscope 2-Plus microscope and imaging system using Zeiss Axiovision version 4.4 software (Carl Zeiss MicroImaging). The average cellular immunofluorescence intensity was quantitated by determining the total intensity of the image, subtracting the calculated cell-free background intensity, and dividing by the number of cells in the image.

Immunoprecipitation of rFPR

Wild-type and rFPR-expressing CHO cells were labeled overnight with Tran-35S-label on 10 cm dishes. Cells were rinsed with PBS and extracted 1 h on ice with 1 ml of buffer containing 0.5% n-dodecyl maltoside, 25 mM Tris-HCl (pH 7.5), 140 mM NaCl, and 2 mM EDTA. Supernatant from a 15 min spin at 12,000 x g was precleared with irrelevant mAb and protein G-Sepharose. Proteins were then immunoprecipitated with NFPR1 and NFPR2 plus protein G-Sepharose. Beads were washed and proteins were released at 55°C for 10 min in Laemmli sample buffer.

Phage display analysis

Phage clones were selected from the J404 random nonapeptide sequence phage display library on NFPR1 or NFPR2-coupled cyanogen bromide-Sepharose, amplified, selected again by plaque replica plating and blotting, and sequenced as described by Burritt et al. (35). Sequences were then visually aligned with one another and by similarity with the FPR sequence (36). Alignments were tabulated in Microsoft Excel. Detailed computational confirmation of epitope mapping of selected phage peptide sequences to the FPR sequence was conducted using a Java version of the program FINDMAP (37), now called EPIALIGN (38), which outputs a frequency histogram showing the number of times each residue of the FPR sequence was computationally aligned with one of the nonapeptide phage display probe peptide residues.

Subcellular fractionation of human neutrophils

Diisopropylfluorophosphate-treated human neutrophils (2.5 x 109) were isolated from two units of blood by the gelatin method (as described above) and disrupted by N2 cavitation in low ionic strength, hypotonic medium (0.34 M sucrose, 10 mM HEPES, 1 mM EDTA, 0.1 mM DTT, 1 mM ATP, 0.1 mM Mg2+, pH 7.4 (with protease inhibitors) as previously described (39). This method produces primed neutrophils devoid of secretory vesicles and plasma membrane fractions without secretory vesicle contamination (31). After 1000 x g centrifugation, the low speed supernatant (28.5 ml) was manually loaded on a 15–60% (w/w) sucrose gradient preformed in a Beckman Ti-14 zonal rotor (Beckman Coulter) according to the manufacturers instructions, overlayed with 46.5 ml of 5% sucrose, and centrifuged at 44,000 rpm for 3.5 h at 4°C. The gradient was pumped from the rotor at 2000 rpm and 50 14-ml fractions were collected manually, protease inhibitors were added and fractions were mixed and evaluated for the distributions of sucrose by refractometer, protein by BCA protein assay, myeloperoxidase using 0.5 mM ABTS as substrate, and alkaline phosphatase using 8 mM p-nitrophenyl phosphate as substrate, or mixed into 4x SDS-PAGE sample buffer for immunoblot densitometry of lactoferrin and FPR.

In-gel and solution digestion of rHis-FPR for MALDI mass spectrometry

In-gel and solution digestion of rHis-FPR and MALDI mass analysis was performed essentially as described by Taylor et al. (40) on the pooled peak fractions eluted from the Ni2+ affinity matrix. Digests were performed on either the Mr ~32 or ~72 kDa bands (Fig. 1, lanes 6, 7, 14, and 15).


Figure 1
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  		FIGURE 1. Purification of rHis-FPR from LPG extracts of Sf9 cell membranes. Sf9 cells infected with baculovirus-bearing recombinant FPR with a hexa-His-N-terminal affinity tag were disrupted by N2 cavitation and membrane fractions prepared. The post nuclear supernatants of the N2 cavitates were successively centrifuged at 100,000 x g to produce the high speed supernatants and pellets after treatment with 10 mM NaOH and subsequent extraction with 1% LPG. The 100,000 x g supernatant from the LPG extraction was then passed over a Ni2+-NTA affinity column and eluted with 300 mM imidazole after washing the column with wash buffer. Membrane proteins from the LPG extract and eluted material from the column were electrophoretically separated by SDS-PAGE and stained with either Coomassie Brilliant Blue (c) or silver nitrate (s) or immunoblotted with an anti-His-tag Ab (i) as a probe for the rHis-FPR. The indicated lanes contained the following material: m.w. markers: 1,10,18; 100KS3, LPG extract: 2(c), 3(s) (illustrates the protein load from which rHis-FPR is affinity purified), 11(i) (illustrates the starting immunoblot signal); material left on Ni2+-NTA column after washes: 4(s),12(i); successive 300 mM imidazole elutions: 5–9(s), 13–17(i). Each lane contains 5 x 105 cell equivalents.

 

   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Preparation of rHis-FPR-enriched Sf9 membranes and purification of rHis-FPR

To produce high levels of rFPR for affinity purification, an N-terminal His-tagged FPR construct was produced in Sf9 cells by infection with rHis-FPR bearing recombinant baculovirus. In brief, the virus was produced by homologous recombination of viral DNA with a pBlueBacHisB-FPR construct. Expression of rHis-FPR was confirmed by immunofluorescence in permeabilized inoculated Sf9 cells using affinity-purified rabbit polyclonal Ab (anti-CTZ) raised against the sequence of the C-terminal 14 amino acid residues of FPR. Compared with nonfluorescent uninfected controls, infection produced very bright specific fluorescence (not shown). Based on this measure and immunoblots using both anti-CTZ Ab and a monoclonal anti-His-tag Ab, it was determined that maximum production of protein occurred 60 h post infection. For purification of rHis-FPR, the infected Sf9 cells were collected, disrupted by N2 cavitation, and the postnuclear supernatant fractionated to generate a crude membrane (100KP1) and alkali-stripped membrane (100KP2) fraction. Recently, LPG (26) was used to recover integral membrane proteins expressed to high levels in Sf9 cells. Lack of sedimentability after 100,000 x g for 1 h was the criterion used for solubilization of receptor. After exposure to 1% LPG for 1 h on ice, >70% of the rHis-FPR was detected in the 100,000 x g supernatants of the detergent extract (100KS3; Fig. 1) and was barely detectable (in overdeveloped blots) in the high speed pellet (not shown), demonstrating successful solubilization of the receptor in LPG.

Affinity purification of LPG-solubilized rHis-FPR was achieved by Ni2+ column chromatography with continuous tumbling of the Ni2+ column matrix and the 100KS3 LPG extract at 4°C overnight. Unbound material in the flow-through and successive washes was collected, and the bound protein was eluted with imidazole. The matrix was sampled and all fractions were analyzed for protein composition by SDS-PAGE, with rHis-FPR content evaluated by immunoblot analysis with anti-CTZ (not shown) and anti-His Ab. Fig. 1 displays the results of the purification for the LPG extracts showing that rHis-FPR, with an apparent relative molecular mass of ~32 kDa, was specifically retained on the Ni2+ matrix and nearly all bound rHis-FPR was eluted from the beads. Additionally, >95% of the protein eluted after washing the matrix (as determined densitometrically) had the same mobility on SDS-PAGE as the immunoblotting species, suggesting that virtually all protein recovered was rHis-FPR. It is noteworthy that alkali stripping of membranes results in a major reduction of masking bands in the range of the monomeric rHis-FPR species resulting from removal of ~2/3 of the membrane proteins (peripherally bound) of the Sf9 membranes with probably little, if any, loss of rHis-FPR (not shown). Based on direct protein measurements using a colorimetric protein assay, typically 200–400 µg of rHis-FPR are produced from 1 liter of Sf9 culture when extracted with LPG. Based on quantitative immunoblotting, the yields represent a 100- to 200-fold enrichment over starting membranes and a recovery of greater than 90% of rHis-FPR protein from the alkali-washed membranes or detergent extract. One reproducible observation in this preparation is the existence of dimers (73 kDa) and, in some instances, higher order multimers (102 kDa) of the 32 kDa band as evidenced by staining with anti-His Ab (Figs. 1 and 2, lane 6) and anti-CTZ peptide Ab (not shown). Because this higher order banding is observed in membranes as well as purified preparations, it is conceivable that they are remnants of higher order oligomerizations of the receptor.


Figure 2
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  		FIGURE 2. Specificity of NFPR1 and NFPR2 for neutrophil and recombinant FPR and for recombinant FPRL1. A, Alkali-stripped membranes from human neutrophils (lanes 2, 8, and 14), wild-type (wt) CHO cells (lanes 3, 9, and 15), rFPR-transfected CHO cells (lanes 4, 10, and 16), uninfected Sf9 cells (lanes 5, 11, and 17), and baculovirus-infected Sf9 cells bearing rHis-FPR (lanes 6, 12, and 18) were used for immunoblotting with anti-His-tag Ab, mAb NFPR1, and mAb NFPR2. Protein load for each of the membrane samples was neutrophil 44.5 µg, CHO 13.5 µg, and Sf9 4.5 µg. Lanes 1, 7, 13, and 19 were loaded with prestained m.w. markers as shown. B, Alkali-stripped membranes from rFPR-expressing CHO cells (lanes 2 and 11), degranulated human neutrophils (lanes 3 and 12), and rFPRL1-transfected TX6 cells (lanes 5–8 and 14–17) and vector control TX31 cells (lanes 9 and 18) were used for immunoblotting with mAb NFPR1 and mAb NFPR2 as shown. Protein load for each of the membrane samples was CHO 0.25 µg, neutrophil 2 µg, TX6 (l to r) 2, 1, 0.5, and 0.25 µg, and TX31 2 µg. Lanes 1, 4, 10, and 13 were loaded with prestained m.w. markers as shown.

 
MALDI-TOF peptide mass mapping of rHis-FPR

To confirm the identity of the purified rHis-FPR, Sf9 cell membrane protein (Mr ~32 and Mr ~72 kDA) purified on the Ni2+ affinity matrix and separated by SDS-PAGE was subject to tryptic digestion (both in-gel and solution) with the resulting samples analyzed by MALDI mass spectrometry. As shown in Table I, a number of observed masses could be assigned to the rHis-FPR sequence, resulting in 36% sequence coverage and the confident identification of the purified protein. It is of interest to note that these MALDI peptide mass mapping studies allowed for the detection of transmembrane helix 4 and suggested that Asn179 (a consensus N-linked glycosylation site) is not modified in Sf9 cells. In addition, the majority of the rHis-FPR C-terminal tail was detected in these studies and demonstrated the potential utility of mass spectrometry for future attempts to identify phosphorylation sites thought to occur in the C-terminal region (see Discussion).


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  		Table I. MALDI peptide mass mapping of recombinant His-FPR

 
Production of monoclonal anti-FPR Abs

Purified rHis-FPR was used as an immunogen to generate mAbs recognizing epitopes of rHis-FPR common to native rFPR and FPR found in primary human cells. NFPR1 and NFPR2 are two mAbs purified from hybridoma supernatants (IgG1 isotype) that were generated from spleen cells of a mouse immunized with rHis-FPR. Both Ab-producing clones produced supernatatants that were found to be negative when screened for binding to intact wild-type and rFPR-expressing CHO cells but positive for binding to permeabilized rFPR-expressing cells (not shown). When screened by immunoblot analysis as described in Materials and Methods, both Abs were positive for a broad band in the range between 40 and 75 kDa relative molecular mass (see below). To confirm that these Abs bind to SDS-denatured FPR expressed in recombinant systems as well as human neutrophils, an immunoblot analysis was performed with membranes obtained from Sf9 cells expressing rHis-FPR, CHO cells expressing rFPR, and membranes obtained from degranulated human neutrophils. Fig. 2 compares the species recognized by NFPR1 and NFPR2 with those recognized by an anti-His-tag Ab. Anti-His-tag Ab primarily recognizes proteins with relative molecular mass 32 and 72 kDa found in rHis-FPR-expressing Sf9 cell membranes but not the non-expressing controls. Anti-CTZ Ab made against a C-terminal peptide produced an identical pattern for Sf9 membranes (not shown) suggesting that the His-tag protein and the CTZ-bearing protein were equivalent. Both NFPR1 and NFPR2 recognize these same species in the infected Sf9 cell extracts, as well as broad species of relative molecular mass 45–75 kDa in both human neutrophil membrane extracts and in membranes prepared from rFPR-expressing CHO cells. Using membranes from fMLF-stimulated, cytochalasin-pretreated human neutrophils (i.e., degranulated neutrophils), NFPR1 also recognized two lower Mr species of ~40 and ~30 kDa. One of these bands may be related chemoattractant receptor FPRL1 (see below). Wild-type non-expressing controls for Sf9 and CHO cells showed negligible nonspecific staining. These results suggest that NFPR1 and NFPR2 recognize several denatured forms of FPR that are generated during SDS-PAGE. Furthermore, this experimental data indicates that human neutrophil FPR, indeed, has similar mobility properties as the rFPR of CHO cells.

Identification of mAb epitopes on FPR

To determine the locus of binding of NFPR1 and NFPR2, a high throughput version of phage display epitope analysis was performed on these two Abs, as we have previously described for other systems (35, 41). Fig. 3 and Fig. 4 show the NFPR1- and NFPR2-selected nonapeptide phage sequences, respectively, aligned according to a visual correspondence of the consensus to specific regions of the predicted cytoplasmic domain of FPR. To confirm this alignment, phage sequences were also examined by the program FINDMAP, which can identify linear and complex epitopes of anti-protein Abs (37, 42). Fig. 5 shows histograms of the most frequently aligned residues of the entire FPR amino acid residue sequence. Both visual and FINDMAP alignments strongly suggest that the NFPR1 Ab recognition site contains residues 305-GQDFRERLI-313 at the juxtamembrane amino-terminal portion of the carboxyl-terminal tail of FPR. Importantly, the FPR amino acid sequence of this region is identically shared by FPRL1. To determine whether NFPR1 also recognizes FPRL1, membranes were prepared from cultures of mouse L fibroblasts stably expressing FPRL1 (25) or vector control. Fig. 2B shows that NFPR1 but not NFPR2 recognizes a single band of approximate relative molecular mass of 40 kDa corresponding to that predicted for FPRL1.


Figure 3
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  		FIGURE 3. Epitope mapping of NFPR1 by phage display analysis. Phage displayed peptide sequences were selected by three successive rounds of affinity retention of clones of the J404 random nonapeptide phage display library on an affinity column of NFPR1 followed by plaque positive identification of Ab-binding clones visualized by replica immunoblot. A linear epitope region of the FPR sequence identified by visual alignment of the phage sequences is shown shaded black. Black and gray shading indicates identity or similarity, respectively, of peptide amino acid residues with FPR sequence residues. The figure is split in half vertically to show all sequences on one page. In the bottom most three rows of the second half of the sequences, the number of identities is scored and the percentages of the visually aligned residues corresponding to the FPR sequence are calculated.

 

Figure 4
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  		FIGURE 4. Epitope mapping of NFPR2 by phage display analysis. Phage displayed peptide sequences were selected by three successive rounds of affinity retention of clones of the J404 random nonapeptide phage display library on an affinity column of NFPR2 followed by plaque positive identification of Ab-binding clones visualized by replica immunoblot. A linear epitope region of the FPR sequence identified by visual alignment of the phage sequences is shown in shaded black. Black and gray shading indicates identity or similarity, respectively, of peptide amino acid residues with FPR sequence residues. In the bottom most three rows, the number of identities is scored and the percentages of the visually aligned residues corresponding to the FPR sequence are calculated.

 

Figure 5
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  		FIGURE 5. FINDMAP epitope mapping of NFPR1 and NFPR2. For FINDMAP identification of epitopes, 103 and 106 nonapeptide sequences were selected from a phage display analysis of NFPR1 (A) and NFPR2 (B) (see Figs. 3 and 4), respectively. These collections were epitope mapped by the computer program FINDMAP (37 ) using the FPR sequence and a substitution matrix allowing for conservative amino acid substitutions. Each nonapeptide sequence was thus aligned with specific residues of the FPR sequence and the best alignments collected used to construct a frequency histogram, which plots the number of times a particular residue in the FPR sequence was identified as part of an alignment. Multiple alignments for any one phage display sequence were weighted inversely according to the number of matches for that particular phage display sequence.

 
Of interest is the observation that the predicted epitope of NFPR2 appears to be more complex than a simple linear epitope, suggesting a strong recognition of the region 337-NSTLPSAEVE-346 of FPR, that shares only 50% identity to the corresponding region of FPRL1. The epitope analysis by FINDMAP on the whole protein sequence reveals other less probable regions shown in Fig. 5B, which include 99-KFVFTIVD-106, 204-IRF-206, and 287-SALAFFN-293, positioned at the extracellular interface of the third predicted transmembrane domain, at the extracellular interface of the fourth transmembrane domain, and in the middle of the seventh predicted transmembrane domain, respectively. Evidence presented below suggests that the Ab binds only intracellular epitopes making these latter possibilities unlikely. Limiting the FINDMAP analysis to the presumed cytoplasmic domains (using sequence alignment with rhodopsin) as the targets of the alignment has identified other regions (see Discussion) that might contribute to this epitope, as well as the epitope identified for NFPR1 (not shown). However, those results await experimental confirmation using Ab imprint analysis with composite synthetic peptides as confirming mimetics of the more complex epitope (35, 42, 43).

Differential activity of NFPR1 and NFPR2

It is of interest that NFPR2 apparently recognizes a region of the carboxyl terminal tail containing strongly implicated S/T phosphorylation sites (17). It could be expected, therefore, that NFPR2 recognition of FPR might be sensitive to the phosphorylation state. To examine this possibility an experiment was performed comparing the relative recognition of rFPR by NFPR1 and NFPR2 in SDS-solubilized whole cell extracts prepared from rFPR-bearing CHO cells pre-exposed to different concentrations of fMLF for different times. Fig. 6A shows that the NFPR1 recognized a broad band of ~60 kDa relative molecular mass that was invariant after fMLF exposure. However, the binding of NFPR2 to a band of the same molecular size was significantly diminished in a time- and concentration-dependent manner. This result would be expected if NFPR2 could not bind its phosphorylated epitope on FPR.


Figure 6
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  		FIGURE 6. Effect of fMLF stimulation and alkaline phosphatase on binding of NFPR1 and NFPR2 to rFPR expressed in CHO cells. A, rFPR-expressing CHO cells were exposed to 0, 0.01, 0.05, 0.1, 0.5, and 1 µM fMLF for 8 min or 1 µM fMLF for 0, 1, 2, 4, 8, or 16 min at 37°C. Cells were solubilized directly in SDS-PAGE sample buffer and then immunoblotted with NFPR1 and NFPR2 after electrophoretic separation and electrotransfer, and developed with chemiluminescence. The concentration and time dependence of the recognition of the prominent, broad ~60-kDa band observed in Fig. 2A, lanes 10 and 16 is shown. B, rFPR expressing CHO cells were exposed to vehicle (-) or 1 µM fMLF (+) for 10 min at 37°C. Alkali-stripped crude membrane fractions were prepared from each population of cells as described in Materials and Methods and aliquots were incubated with 10 U of SAP at 37°C for 0, 0.2, 0.5, 1, 2, or 16 h in 0.3 M Tris-HCl, pH 7.3, 0.3 M NaCl or with SAP and phosphatase inhibitor mixture for 2 h. To remove enzyme these membranes were diluted 20-fold and pelleted at 100,000 x g for 30 min before electrophoresis and immunoblotting with NFPR1 and NFPR2.

 
To confirm that the rFPR modification in the NFPR2 epitope is indeed phosphorylation, membranes from stimulated and unstimulated CHO cells were exposed to shrimp alkaline phosphatase (SAP) (31) at 37°C, washed by ultracentrifugation, and then immunoblotted and quantitated densitometrically. Fig. 6B shows that the density of the rFPR band probed with NFPR2 is recovered at progressively increasing times of alkaline phosphatase treatment between 0 and 16 h. When probed with NFPR1, immunolabeling density is relatively insensitive to SAP treatment. The measured density after 16 h of incubation with SAP is ~50 to 80% of the unstimulated control CHO cell membranes untreated with SAP. If phosphatase inhibitor mixture is included in the reaction, this recovery does not occur (Fig. 6B). Similar changes, increasing the labeling density of the ~60 kDa band by 70 ± 20% (n = 6), could be observed when membranes of stimulated degranulated human neutrophils shown in Fig. 2A, lane 14 were exposed to SAP for 2 h at 37°C (not shown). Although neutrophils have endogenous membrane-bound alkaline phosphatase activity, it is topologically sequestered from the intracellular faces of neutrophil plasma membrane or granule membrane, thus minimizing its significance for FPR. Together, these results suggest that NFPR2 but not NFPR1 recognition of denatured, electroblotted rFPR and FPR is sensitive to the phosphorylation state of the receptor.

NFPR2 recognizes an epitope on detergent-solubilized Tran35S-labeled rFPR

Because one of the methods of mAb screening indicated that both Abs in the hybridoma supernatants could specifically bind to rFPR-expressing CHO cells and not wild-type CHO cells, it seemed possible that Abs would be able to bind to native or at least partially denatured FPR. To demonstrate that such was the case, we examined the ability of both Abs to immunoprecipitate rFPR from nonionic detergent extracts. CHO, wild-type, and rFPR-expressing cells, were grown in the presence of Tran35S label, extracted in 0.5% n-dodecyl maltoside, clarified by centrifugation at 12,000 x g and precleared by exposure with irrelevant mAb and protein G-Sepharose. Proteins were then immunoprecipitated with NFPR1 and NFPR2 bound to protein G-Sepharose, washed and extracted in sample buffer for SDS-PAGE. PhosphorImager analysis and conventional autoradiography were performed on dried SDS-PAGE gels. As shown in Fig. 7, only NFPR2 was capable of specifically immunoprecipitating a radioactive broad band of mobility equivalent to the Mr ~60 kDa protein in the rFPR-expressing cells, suggesting that CHO-FPR epitope for NFPR1 may not be accessible by this Ab when solubilized in dodecyl maltoside.


Figure 7
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  		FIGURE 7. Immunoprecipitation of Tran35S-labeled FPR by NFPR2 but not NFPR1. Wild-type (lanes 1 and 3) and rFPR-expressing CHO cells (lanes 2 and 4) were labeled overnight with Tran35S-label, rinsed, extracted with dodecyl maltoside and then immunoprecipitated with NFPR1 (lanes 1 and 2) and NFPR2 (lanes 3 and 4) coupled to protein G-Sepharose. Beads were washed and proteins were released at 55°C for 10 min in Laemmli sample buffer. PhosphorImager analysis (left) and conventional autoradiography (right) performed on the same dried SDS-PAGE gel is shown.

 
Immunolocalization of FPR in human neutrophil subcellular fractions

To confirm the subcellular localization of FPR in human neutrophils, immunoblot analysis was performed on fractions collected from isopycnic sucrose gradient separations of homogenates of human neutrophils prepared by the gelatin method (44). The location of the major organelles in the gradient were identified with alkaline phosphatase ±0.1% Triton X-100, lactoferrin, and myeloperoxidase activity serving as markers for light plasma membrane (peak at 32% sucrose), specific granules (peak at 42% sucrose) and azurophil granules (peak at 48% sucrose), respectively (Fig. 8) (10, 34). There was no detectable alkaline phosphatase, myeloperoxidase or lactoferrin in the cytosolic fractions. NFPR1 and NFPR2 were used to measure the distributions of the immunoblotted bands in gradient fractions. Fig. 8 shows the distributions of the ~60 kDa species recognized by both Abs to be virtually identical to one another and to alkaline phosphatase in the plasma membrane region. It should be noted that primed neutrophils purified by the gelatin method contain no secretory vesicles (34, 44) and all alkaline phosphatase is expressed on the cell surface with no latent activity uncovered in the presence of Triton X-100. Thus the distribution of the ~60 kDa species suggests an 85% surface localization of the band as detected by both Abs. Of interest is the significantly reduced level of the ~60 kDa species in the lactoferrin-enriched fractions also observed in these distributions (Fig. 8C). These profiles confirm previous observations that in primed neutrophils most FPR is found in the plasma membrane and that the intracellular granule stores are effectively depleted (45, 46). Interestingly, the 40 kDa band, recognized by NFPR1 and suggestive of the distribution of FPRL1, appeared evenly distributed between the two fractions (Fig. 8, C and D). The 30-kDa band also recognized by NFPR1, demonstrated a peak in the heavy plasma membrane fraction (not shown), corresponding to a cytoskeleton-enriched plasma membrane domain (44). All three bands are apparent when cells are degranulated as is shown in Fig. 2, lane 8. This unidentified ~30-kDa band appears to be variably present in the membrane, and it may arise from an association of one of the strong doublet bands observed in the cytosolic fraction at approximately Mr ~30 kDa, when immunostaining with this Ab. If the cytosol is not subjected to denaturing conditions of SDS-PAGE, it is not recognized in a dot blot analysis using similar and much higher loads (not shown). Results were equivalent in the high-resolution gradient shown in Fig. 8 as well as in lower resolution gradients centrifuged in smaller swinging bucket rotors (not shown).


Figure 8
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  		FIGURE 8. Subcellular localization of human neutrophil FPR by isopycnic centrifugation of N2 cavitates from unstimulated human neutrophils. Primed human neutrophils were prepared from whole blood using the 1 x g sedimentation gelatin method (34 ). The postnuclear supernatant fractions of N2 cavitates were separated by isopycnic sedimentation in linear sucrose gradients as described in Materials and Methods. A, Total protein and sucrose content of fractions are plotted against fraction number as a reference. B, Total protein, surface and total alkaline phosphatase (AP) activity in the presence (+) and absence (—) of 0.1% Triton X-100, lactoferrin (LF), and myeloperoxidase (MPO) activities were measured and plotted as a function of fraction sucrose percentage to identify the fractions enriched in cytosol (total protein), plasma membrane (AP) specific granules (LF), and azurophil granules (MPO), respectively. C, NFPR1 and NFPR2 immunoblotting activities are plotted as a function of percent sucrose of each fraction from 20 to 60% sucrose. Relative densities are shown for multiple bands observed in Fig. 2 that were developed by NFPR1 at Mr ~60 kDa (NFPR1–60), ~40 kDa (NFPR1–40). Only a single band was observed for NFPR2 at ~60 kDa. No FPR was detected in the cytosol fraction. As a reference, alkaline phosphatase activity and lactoferrin activity shown in B are plotted as dotted and dashed/dotted lines, respectively. D, Peak alkaline phosphatase-enriched (PM) and lactoferrin-enriched (SG) fractions were immunoblotted with NFPR1 and NFPR2 as described in Materials and Methods and developed using chemiluminescence. Relative molecular mass markers are in kDa as shown.

 
Immunofluorescence detection of FPR

To confirm FPR localization in human neutrophils as implied by the sucrose gradient sedimentation results and sensitivity of NFPR2 to stimulation of neutrophils by fMLF, indirect immunofluorescence analysis was performed by flow cytometry as well as fluorescence microscopy. The first part of this analysis used methanol fixed and permeabilized neutrophils that had been exposed to 1 µM fMLF or vehicle. Fig. 9A shows a flow cytometry analysis of the degree of permeabilized neutrophil labeling by these Abs. The geometric mean fluorescence level (–/+fMLF) for each category was as follows: NFPR1 (57/49), NFPR2 (42/26), K16 (11/11), diluent (7/7). These results show first that both Abs bind specifically to fMLF-stimulated and unstimulated, fixed and permeabilized neutrophils. Secondly, the results also show that NFPR1 is insensitive to fMLF stimulation whereas NFPR2 is sensitive to cellular activation state, binding less efficiently to stimulated cells. The right panel shows a control comparing the labeling intensity of K16 to fixed and permeabilized cells exposed to diluent instead of primary Ab and shows, at most, a 15% reduction when primary Abs were withheld. It was also found that NFPR1 efficiently binds methanol- but not formalin-fixed cells whereas NFPR2 binds more efficiently to formalin-fixed cells (not shown). Intact, nonpermeabilized CHO cells showed no binding of Ab to the extracellular face of the plasma membrane, so nonpermeabilized neutrophils were not examined further.


Figure 9
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  		FIGURE 9. Flow cytometry and immunofluorescence microscopy of unstimulated and fMLF-stimulated human neutrophils. Human neutrophils were prepared by 1 x g sedimentation in 2% gelatin, treated with diisopropylfluorophosphate, and warmed to 37°C. Half of the cells were exposed to vehicle (—) and the other half to 1 µM fMLF for 5 min (+). A, For flow cytometry analysis the cells were fixed, permeabilized, washed, and labeled as described in Materials and Methods using 10 µg/ml NFPR1, NFPR2, K16 (an isotype matched irrelevant control), and diluent only. These cells were washed, labeled with an Alexa Fluor 488-labeled secondary goat anti-mouse polyclonal Ab, and washed again. Analysis by flow cytometry used 488 nm excitation and a 520 nm emission cutoff. The geometric mean fluorescence level (±fMLF) for each category was as follows: NFPR1 (57/49), NFPR2 (42/26), K16 (11/11), diluent (7/7). B, Live cells treated as described above were sedimented onto glass slides with a cytospin, followed by cold methanol fixation. Ab binding and immunofluorescence staining, as described in Materials and Methods, were conducted with three intervening wash steps following each labeling step. The microscopy results show composite immunofluorescence images of 12 to 13 cells for unstimulated neutrophils (a, b, c) and fMLF-stimulated neutrophils (d, e, f), labeled with NFPR1 (a, d), NFPR2 (b, e), and control K16 (c, f). The average relative intensities of cells in a–f, based on integrated intensities of the fields containing cells minus background, are 6.9, 5.1, 1.6, 7.3, 3.1, 1.9, respectively. Panel edge = 50 µm.

 
To further confirm FPR localization in neutrophils, the NFPR Abs were also used to examine receptor labeling by immunofluorescence microscopy. Gelatin-prepared (primed) neutrophils, exposed to 1 µM fMLF or vehicle, were sedimented directly onto glass slides by cytospin centrifugation followed by fixation with methanol and labeling with either NFPR1, NFPR2, or control irrelevant Ab K16. Fig. 9, a and b, show NFPR1 and NFPR2 immunofluorescence labeling, respectively, of unstimulated flattened neutrophils. Fig. 9, d and e, show the corresponding labeling of neutrophils stimulated with 1 µM fMLF at 37°C and confirm that the intensity of labeling by NFPR2 is reduced in the stimulated population. Fig. 9, c and f, show negligible labeling by the irrelevant control Ab K16. It should be noted that formalin fixation seems to preserve immunofluorescence staining by NFPR2 but is negative with NFPR1 (not shown).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
The N-formyl peptide receptor has long served as a model for the signal transduction operating in the activation of human phagocytes (3, 47). The occupancy of FPR by formylated peptides induces cell polarization, intracellular calcium release, phosphatidyl inositol metabolism, phospholipase C and D activation, and the activation of a complex cascade of intracellular kinases (3, 48). FPR-effector responses, that result from some or all of these changes, include morphological polarization, pseudopodial orientation, adhesion, migration up concentration gradients, degranulation, secretion, and superoxide generation (49). These processes are manifestations of the host defensive machinery, without which the host would be unable to survive infection. An essential step in the defense process is the delivery of massive numbers of neutrophils to sites of infection or injury. The accumulation of these cells is so great that they often overwhelm tissue cells, causing the hallmark pathology observed in nearly every type of inflammatory reaction examined by pathologists (50). One interesting aspect of neutrophil accumulation is how these inflammatory cells are able to maintain migration over gradients of chemoattractants that range by several orders of magnitude. Such high exposures suggest that surface chemoattractant receptors would be saturated and insensitive to differential concentration of chemoattractant spanning the dimension of the chemotactic cell (52). Hence the adaptive regulatory processes affecting receptor affinity, surface distribution, re-expression and internalization, as well as coupling efficiency of inflammatory receptors in isolated primary human neutrophils and those in inflammatory loci need to be understood.

To better address these issues, we have generated two novel mAbs that recognize FPR. To accomplish this, we purified a His-tagged FPR expressed in Sf9 cells and used it as an immunogen for the production of mAbs in mouse spleen cell-derived hybridomas. These Abs were epitope-mapped using phage display Ab imprint analysis, which suggested that they recognized the cytoplasmic tail of the protein in two different regions. One of these regions, 337-NSTLPSAEVE-346 recognized by NFPR2, includes S/T phosphorylation sites, whereas the other, 305-GQDFRERLI-313 recognized by NFPR1, does not. A more intensive computer analysis limiting the alignments to only the cytoplasmic regions using the program FINDMAP has suggested that lesser contributions to the epitope of NFPR2 may include the region 323-ALTE-326, 308-DFRE-311, 121-LD-122, and 62-ISYL-65; however, the relative contribution of these regions needs to be confirmed by Ab imprint analysis (35, 42, 43). 305-GQDFRERLI-313 of FPR is also identical in FPRL1, a low-affinity formyl peptide receptor (25, 52) having a wider variety of specific higher affinity ligands (1), including lipoxin A4 (53), which may play an important role in disease (52) and the resolution of inflammation (54). These findings suggest that NFPR1 and NFPR2 might be useful as probes of FPR and FPRL1.

Both of these anti-FPR mAbs were able to specifically bind recombinant FPR in the Sf9 and CHO cell expression systems, whereas NFPR1 also bound recombinant FPRL1 specifically expressed in mouse L cell fibroblasts. These results along with the phage display epitope analyses for these two Abs strongly suggest that the corresponding Mr ~60- and 40-kDa species observed in neutrophils also is indicative of these two receptors. Interestingly, these two species localize differently in the plasma membrane and intracellular stores. The ~60-kDa species demonstrates a closely correlated cosedimentation with alkaline phosphatase (~85% of the total) and a minority (~15% of the total) cosedimentation with the lactoferrin-enriched, specific granule/gelatinase granule-containing fractions (55, 56), consistent with our previous determinations of the subcellular localization of tritiated fMLF binding (39), iodinated formyl peptide photoaffinity labeling (10), and studies performed in other laboratories (46) on primed neutrophils. The Mr ~40-kDa band also demonstrates similar loci but with approximately equal amounts of 40-kDa species in each. Because the fractionation was performed on primed cells devoid of secretory granules it seems probable that the two receptor types are differentially regulated.

In addition to the Mr ~40- and 60-kDa bands, NFPR1 also binds to a 30-kDa species that is observed in the membranes of degranulated and unstimulated neutrophils (Figs. 2, and 8D, respectively), in the membranes and granules of unstimulated neutrophils (Fig. 8D), as well as the cytosolic fractions. This added cross reactivity limits the molecular interpretation of NFPR1 to immunoblotting data where bands have been separated by SDS-PAGE. It precludes unambiguous molecular interpretation of the interesting NFPR1-specific immunofluorescence distributions observed in Fig. 9, a and d. However, it should be noted that the cytosolic Mr ~30 kDa does not appear to be bound by the Ab unless it is denatured and separated by SDS-PAGE, supporting the interpretation that the immunofluorescence distributions do not correspond to the cytosolic species.

NFPR2 immunofluorescence labeling in neutrophils, however, appears to be more interpretable. Only a single species of Mr ~60 kDa appears to be labeled in immunoblots of neutrophil membranes after SDS-PAGE separation. This band demonstrates sensitivity to the fMLF stimulation history of the neutrophils (J. Stie and A. Jesaitis, unpublished data) and an alkaline phosphatase-mediated recovery of binding in immunoblots of membranes obtained from fMLF-stimulated cytochalasin B-pretreated neutrophils shown in Fig. 2. These observations parallel those made using rFPR-expressing CHO cell fractions. Considering also that NFPR2 was raised against a FPR polymorphic form that is expressed in the human population infrequently (57), it probably exhibits a somewhat different cross reactivity with the more abundant polymorphic FPR form, expressing alanine 346 instead of the negatively charged glutamic acid 346. However, because NFPR2 recognition of its epitope is substantially diminished by stimulation FPR-bearing cells, our results are consistent with fMLF-dependent phosphorylation of any of the potential S/T phosphorylation sites T336, S338, T339, S342 in or adjacent to the NFPR2 epitope. This inference is supported by the results of Prossnitz and coworkers (5, 17, 58) showing that the site-directed mutagenesis of S336, T339 may affect ligand-induced receptor processing, in ways that are consistent with receptor phosphorylation. We therefore conclude that 1) the Mr ~60-kDa band recognized by NFPR1 and NFPR2 in neutrophil membranes is FPR; 2) that the NFPR2 immunofluorescence distribution in fixed unstimulated neutrophils probably represents that of FPR; and that 3) the loss of NFPR2 labeling of neutrophils with fMLF stimulation probably reflects its C-terminal tail phosphorylation and/or sequestration.

A definitive demonstration of phosphorylation of these residues in either system by mass spectrometry or some other direct method must be awaited. In preliminary efforts at MALDI peptide mass mapping of human neutrophil sucrose density gradient fractions showing peak FPR distribution (see Fig. 8), SDS-PAGE gel slices (loading estimated to contain ~40 pmol of total FPR) spanning the ~27- to 75-kDa range were removed from unstained gels and subject to tryptic digestion. The resulting spectra were complex and demonstrated a large number of abundant peaks (indicative of multiple proteins in each gel slice), with four peaks potentially corresponding to human FPR (residues 86–99, 191–201, 231–241, 312–322) using theoretical tryptic digests of all known isoforms (R. Taylor and A. Jesaitis, unpublished). Current studies are focused on developing methods for the further enrichment of human neutrophil FPR to facilitate direct mass analysis of FPR post-translational modification by both MALDI and LC-MS/MS.

The observed subcellular distributions of the Mr ~60- and 40-kDa bands detected by NFPR1 and NFPR2 suggest that FPR and FPRL1 are differentially distributed in these fractions. This result may mean that most of FPR is on the surface of primed neutrophils and FPRL1 is equally abundant on the surface and in intracellular granule stores. These observations are consistent with those reported by Bylund et al. (55) who showed that LPS induced priming up-regulated FPRL1 to the plasma membrane from specific/gelatinase granule stores. Because the affinity of FPRL1 for formyl peptides is estimated to be two to three orders of magnitude lower than that of FPR (25), this observation supports their possible coordinate role in chemotaxis of neutrophils over broader ranges of formyl peptides than suggested by the individual reported affinities and for the adaptive response of neutrophils to formyl peptides (59). Indeed, later delivery of FPRL1 to the surface of neutrophils might also play a role in the regulation of chemotaxis at high chemoattractant concentrations or in the resolution of inflammation by the lipoxin A4 activity of FPRL1 (53) either intracellularly or at the cell surface.

The NFPR Abs reported in our work should be useful for examining FPR localization in human neutrophils during various states of activation and polarization, especially if coupled with subcellular fractionation studies. These Abs provide a significant leap in sensitivity and reduction of nonspecific background that, to our knowledge, has not been achievable with rabbit or mAbs specific for FPR.


   Acknowledgments
 
We thank Dr. James Burritt of the Department of Microbiology for help with immunofluorescence microscopy and Dr. Bruce Granger for help with the flow cytometry. Special thanks also to Dr. Richard Ye who provided the FPRL1-expressing mouse L fibroblasts.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
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 U.S. Public Health Service Grants 2R01-AI26711, 2R01-AI22735 (to A.J.J.), R01-AI51726 (to H.M.M.), and the American Heart Association Scientist Development Grant 06302S3N (to R.M.T.). Back

2 Address correspondence and reprint requests to Dr. Algirdas Jesaitis, Montana State University, 109 Lewis Hall, Bozeman, MT 59717. E-mail address: umbaj{at}montana.edu Back

3 Abbreviations used in this paper: FPR, N-formyl peptide receptor; GPCR, G protein-coupled receptor; LPG, lysophosphatidyl glycerol; fMLF, f-Met-Leu-Phe; CNBr, cyanogen bromide; CHO, Chinese hamster ovary; DPBS, Dulbecco’s PBS; SAP, shrimp alkaline phosphatase; CTZ, c-terminal 337-FPR-350 peptide. Back

Received for publication October 23, 2006. Accepted for publication May 22, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

  1. Fu, H., J. Karlsson, J. Bylund, C. Movitz, A. Karlsson, C. Dahlgren. 2006. Ligand recognition and activation of formyl peptide receptors in neutrophils. J. Leukocyte Biol. 79: 247-256. [Free Full Text]
  2. Prossnitz, E. R., R. D. Ye. 1997. The N-formyl peptide receptor: a model for the study of chemoattractant receptor structure and function. Pharmacol. Ther. 74: 73-102. [Medline]
  3. Mills, J. S., H. M. Miettinen, A. J. Jesaitis. 1999. The N-formyl peptide receptor: structure, signaling, and disease. C. N. Serhan, and P. A. Ward, eds. Molecular and Cellular Basis of Inflammation 215-246. Humana Press, Totowa.
  4. Huang, C.-K.. 1987. Partial purification and characterization of formylpeptide receptor from rabbit peritoneal neutrophils. J. Leukocyte Biol. 41: 63-69. [Abstract]
  5. Prossnitz, E. R., C. M. Kim, J. L. Benovic, R. D. Ye. 1995. Phosphorylation of the N-formyl peptide receptor carboxyl terminus by the G protein-coupled receptor kinase, GRK2. J. Biol. Chem. 270: 1130-1137. [Abstract/Free Full Text]
  6. Diviani, D., A. L. Lattion, S. Cotecchia. 1997. Characterization of the phosphorylation sites involved in G protein-coupled receptor kinase- and protein kinase C-mediated desensitization of the {alpha}1B-adrenergic receptor. J. Biol. Chem. 272: 28712-28719. [Abstract/Free Full Text]
  7. Zhang, J., S. S. Ferguson, L. S. Barak, S. R. Bodduluri, S. A. Laporte, P. Y. Law, M. G. Caron. 1998. Role for G protein-coupled receptor kinase in agonist-specific regulation of mu-opioid receptor responsiveness. Proc. Natl. Acad. Sci. USA 95: 7157-7162. [Abstract/Free Full Text]
  8. Vaughan, R. A., P. N. Devreotes. 1988. Ligand-induced phosphorylation of the cAMP receptor from Dictyostelium discoideum. J. Biol. Chem. 263: 14538-14543. [Abstract/Free Full Text]
  9. Jesaitis, A. J., K.-N. Klotz. 1993. Cytoskeletal regulation of chemotactic receptors: molecular complexation of N-formyl peptide receptors with G proteins and actin. Eur. J. Hematol. 51: 288-293. [Medline]
  10. Jesaitis, A. J., J. O. Tolley, G. M. Bokoch, R. A. Allen. 1989. Regulation of chemoattractant receptor interaction with transducing proteins by organizational control in the plasma membrane of human neutrophils. J. Cell Biol. 109: 2783-2790. [Abstract/Free Full Text]
  11. Zigmond, S. H., S. J. Sullivan, D. A. Lauffenburger. 1982. Kinetic analysis of chemotactic receptor modulations. J. Cell Biol. 92: 34-43. [Abstract/Free Full Text]
  12. Key, T. A., T. D. Foutz, V. V. Gurevich, L. A. Sklar, E. R. Prossnitz. 2003. N-formyl peptide receptor phosphorylation domains differentially regulate arrestin and agonist affinity. J. Biol. Chem. 278: 4041-4047. [Abstract/Free Full Text]
  13. Gripentrog, J. M., A. J. Jesaitis, H. M. Miettinen. 2000. A single amino acid substitution (N297A) in the conserved NPXXY sequence of the human N-formyl peptide receptor results in inhibition of desensitization and endocytosis, and a dose-dependent shift in p42/44 mitogen-activated protein kinase activation and chemotaxis. Biochem. J. 352: (Pt 2):399-407. [Medline]
  14. Bockaert, J., J. P. Pin. 1999. Molecular tinkering of G protein-coupled receptors: an evolutionary success. EMBO J. 18: 1723-1729. [Medline]
  15. Pao, C. S., J. L. Benovic. 2002. Phosphorylation-independent desensitization of G protein-coupled receptors?. Sci. STKE 2002: E42
  16. Tardif, M., L. Mery, L. Brouchon, F. Boulay. 1993. Agonist-dependent phosphorylation of N-formyl peptide and activation peptide from the fifth component of C (C5a) chemoattractant receptors in differentiated HL60 cells. J. Immunol. 150: 3534-3545. [Abstract]
  17. Potter, R. M., D. C. Maestas, D. F. Cimino, E. R. Prossnitz. 2006. Regulation of N-formyl peptide receptor signaling and trafficking by individual carboxyl-terminal serine and threonine residues. J. Immunol. 176: 5418-5425. [Abstract/Free Full Text]
  18. Prossnitz, E. R., O. Quehenberger, C. G. Cochrane, R. D. Ye. 1991. Transmembrane signalling by the N-formyl peptide receptor in stably transfected fibroblasts. Biochem. Biophys. Res. Commun. 179: 471-476. [Medline]
  19. Felgner, P. L., T. R. Gadek, M. Holm, R. Roman, H. W. Chan, M. Wenz, J. P. Northrop, G. M. Ringold, M. Danielsen. 1987. Lipofection: a highly efficient, lipid-mediated DNA-transfection procedure. Proc. Natl. Acad. Sci. USA 84: 7413-7417. [Abstract/Free Full Text]
  20. Miettinen, H. M., J. S. Mills, J. M. Gripentrog, E. A. Dratz, B. L. Granger, A. J. Jesaitis. 1997. The ligand binding domain of the formyl peptide receptor maps in the transmembrane region. J. Immunol. 159: 4045-4054. [Abstract]
  21. Henson, P. M., Z. G. Oades. 1975. Stimulation of human neutrophils by soluble and insoluble immunoglobulin aggregates. J. Clin. Invest. 56: 1053-1061. [Medline]
  22. Summers, M. D., G. E. Smith. 1987. A Manual of Methods for Baculovirus Vectors and Insect Cell Culture Procedures. Texas Agricultural Experiment Station Bulletin No. 1555 Texas Agricultural Experiment Station, College Station, TX.
  23. Parkos, C. A., R. A. Allen, C. G. Cochrane, A. J. Jesaitis. 1987. Purified cytochrome b from human granulocyte plasma membrane is composed of two polypeptides with relative molecular weights of 91,000 and 22,000. J. Clin. Invest. 80: 732-742. [Medline]
  24. Jesaitis, A. J., R. W. Erickson, K.-N. Klotz, R. K. Bommakanti, D. W. Siemsen. 1993. Functional molecular complexes of human N-formyl chemoattractant receptors and actin. J. Immunol. 151: 5653-5665. [Abstract]
  25. Ye, R. D., S. L. Cavanagh, O. Quehenberger, E. R. Prossnitz, C. G. Cochrane. 1992. Isolation of a cDNA that encodes a novel granulocyte N-formyl peptide receptor. Biochem. Biophys. Res. Commun. 184: 582-589. [Medline]
  26. Huang, P., Q. Liu, G. A. Scarborough. 1998. Lysophosphatidylglycerol: a novel effective detergent for solubilizing and purifying the cystic fibrosis transmembrane conductance regulator. Anal. Biochem. 259: 89-97. [Medline]
  27. Laemmli, U. K.. 1970. Cleavage of structural protein during assembly of the head of bacteriophage T4. Nature 222: 680-685.
  28. Wuestehube, L. J., D. W. Speicher, A. Shariff, E. J. Luna. 1991. F-actin affinity chromatography of detergent-solubilized plasma membranes: purification and initial characterization of ponticulin from Dictyostelium discoideum. Methods Enzymol. 196: 47-65. [Medline]
  29. Wray, W., T. Boulikas, V. Wray, R. J. Hancock. 1981. Silver staining of protein in polyacrylamide gels. Anal. Biochem. 118: 197-203. [Medline]
  30. Suvorova, E. S., J. M. Gripentrog, H. M. Miettinen. 2005. Different endocytosis pathways of the C5a receptor and the N-formyl peptide receptor. Traffic 6: 100-115. [Medline]
  31. Khosravi, R., R. Maya, T. Gottlieb, M. Oren, Y. Shiloh, D. Shkedy. 1999. Rapid ATM-dependent phosphorylation of MDM2 precedes p53 accumulation in response to DNA damage. Proc. Natl. Acad. Sci. USA 96: 14973-14977. [Abstract/Free Full Text]
  32. Parkos, C. A., S. P. Colgan, T. W. Liang, A. Nusrat, A. E. Bacarra, D. K. Carnes, J. L. Madara. 1996. CD47 mediates post-adhesive events required for neutrophil migration across polarized intestinal epithelia. J. Cell Biol. 132: 437-450. [Abstract/Free Full Text]
  33. Zola, H., M. W. Brooks. Techniques for the production and characterization of monoclonal hybridoma antibodies. J. G. R. Hurrell, ed. Monoclonal Hybridoma Antibodies: Techniques and Applications 1 CRC Press, Boca Raton.
  34. Stie, J., A. J. Jesaitis. 2004. Preparation of secretory vesicle-free plasma membranes by isopycnic sucrose density gradient fractionation of neutrophils purified by the gelatin method. Cytotechnology 46: 109-122. [Medline]
  35. Burritt, J. B., S. C. Busse, D. Gizachew, D. W. Siemsen, M. T. Quinn, C. W. Bond, E. A. Dratz, A. J. Jesaitis. 1998. Antibody imprint of a membrane protein surface: Phagocyte flavocytochrome b. J. Biol. Chem. 273: 24847-24852. [Abstract/Free Full Text]
  36. Boulay, F., M. Tardif, L. Brouchon, P. Vignais. 1990. The human N-formylpeptide receptor: characterization of two cDNA isolates and evidence for a new subfamily of G-protein-coupled receptors. Biochemistry 29: 11123-11133. [Medline]
  37. Mumey, B. M., B. W. Bailey, B. Kirkpatrick, A. J. Jesaitis, T. Angel, E. A. Dratz. 2003. A new method for mapping discontinuous antibody epitopes to reveal structural features of proteins. J. Comput. Biol. 10: 555-567. [Medline]
  38. Mumey, B., N. Ohler, T. Angel, A. Jesaitis, E. Dratz. 2006. Filtering epitope alignments to improve protein surface prediction. In Lecture Notes in Computer Science Vol 4331/2006. Frontiers of High Performance Computing and Networking-ISPA 2006 Workshops 648-657. Springer, Berlin/Heidelberg.
  39. Jesaitis, A. J., J. R. Naemura, R. G. Painter, L. A. Sklar, C. G. Cochrane. 1982. Intracellular localization of N-formyl chemotactic receptor and Mg2+ dependent ATPase in human granulocytes. Biochim. Biophys. Acta 719: 556-568. [Medline]
  40. Taylor, R. M., D. Baniulis, J. B. Burritt, J. Gripentrog, C. I. Lord, M. Riesselman, W. Maaty, B. P. Bothner, T. E. Angel, E. A. Dratz, H. L. Malech, A. J. Jesaitis. 2006. Analysis of Human Phagocyte Flavocytochrome b by Mass Spectrometry. J. Biol. Chem. 281: 37045-37056. [Abstract/Free Full Text]
  41. Burritt, J. B., M. T. Quinn, M. A. Jutila, C. W. Bond, A. J. Jesaitis. 1995. Topological mapping of neutrophil cytochrome b epitopes with phage-display libraries. J. Biol. Chem. 270: 16974-16980. [Abstract/Free Full Text]
  42. Bailey, B. W., B. Mumey, P. A. Hargrave, A. Arendt, O. P. Ernst, K. P. Hofmann, P. R. Callis, J. B. Burritt, A. J. Jesaitis, E. A. Dratz. 2003. Constraints on the conformation of the cytoplasmic face of dark-adapted and light-excited rhodopsin inferred from antirhodopsin antibody imprints. Protein Sci. 12: 2453-2475. [Medline]
  43. Jesaitis, A. J., D. Gizachew, E. A. Dratz, D. W. Siemsen, K. C. Stone, J. B. Burritt. 1999. Actin surface structure revealed by antibody imprints: evaluation of phage-display analysis of anti-actin antibodies. Protein Sci. 8: 760-770. [Medline]
  44. Stie, J., A. J. Jesaitis. 2007. Reorganization of the human neutrophil plasma membrane is associated with functional priming: implications for neutrophil preparations. J. Leukocyte Biol. 81: 672-685. [Abstract/Free Full Text]
  45. Jesaitis, A. J., G. M. Bokoch, J. O. Tolley, R. A. Allen. 1988. Lateral segregation of occupied chemotactic receptors into actin and fodrin-rich plasma membrane microdomains depleted in guanyl nucleotide proteins. J. Cell Biol. 107: 921-928. [Abstract/Free Full Text]
  46. Sengelov, H., F. Boulay, L. Kjeldsen, N. Borregaard. 1994. Subcellular localization and translocation of the receptor for N-formylmethionyl-leucyl-phenylalanine in human neutrophils. Biochem. J. 299: 473-479. [Medline]
  47. Williams, L. T., R. Snyderman, M. C. Pike, R. J. Lefkowitz. 1977. Specific receptor sites for chemotactic peptides on human polymorphonuclear leukocytes. Proc. Natl. Acad. Sci. USA 74: 1204-1208. [Abstract/Free Full Text]
  48. Prossnitz, E. R., R. D. Ye. 1997. The N-formyl peptide receptor: a model for the study of chemoattractant receptor structure and function. Pharmacol. Ther. 74: 73-102. [Medline]
  49. Snyderman, R., E. J. Goetzl. 1981. Molecular and cellular mechanisms of leukocyte chemotaxis. Science 213: 830-835. [Abstract/Free Full Text]
  50. Gallin, J. I., I. M. Goldstein, R. Snyderman. 1988. Inflammation: Basic Principles and Clinical Correlates 1 Raven Press, New York.
  51. Painter, R. G., L. A. Sklar, A. J. Jesaitis, M. Schmitt, C. G. Cochrane. 1984. Activation of neutrophils by N-formyl chemotactic peptides. Fed. Proc. 43: 2737-2742. [Medline]
  52. Su, S. B., W. Gong, J. L. Gao, W. Shen, P. M. Murphy, J. J. Oppenheim, J. M. Wang. 1999. A seven-transmembrane, G protein-coupled receptor, FPRL1, mediates the chemotactic activity of serum amyloid A for human phagocytic cells. J. Exp. Med. 189: 395-402. [Abstract/Free Full Text]
  53. Chiang, N., I. M. Fierro, K. Gronert, C. N. Serhan. 2000. Activation of lipoxin A(4) receptors by aspirin-triggered lipoxins and select peptides evokes ligand-specific responses in inflammation. J. Exp. Med. 191: 1197-1208. [Abstract/Free Full Text]
  54. Serhan, C. N., B. Levy. 2003. Novel pathways and endogenous mediators in anti-inflammation and resolution. Chem. Immunol. Allergy 83: 115-145. [Medline]
  55. Bylund, J., A. Karlsson, F. Boulay, C. Dahlgren. 2002. Lipopolysaccharide-induced granule mobilization and priming of the neutrophil response to Helicobacter pylori peptide Hp(2–20), which activates formyl peptide receptor-like 1. Infect. Immun. 70: 2908-2914. [Abstract/Free Full Text]
  56. Dewald, B., U. Bretz, M. Baggiolini. 1982. Release of gelatinase from a novel secretory compartment of human neutrophils. J. Clin. Invest. 70: 518-525. [Medline]
  57. Sahagun-Ruiz, A., J. S. Colla, J. Juhn, J. L. Gao, P. M. Murphy, D. H. McDermott. 2001. Contrasting evolution of the human leukocyte N-formylpeptide receptor subtypes FPR and FPRL1R. Genes Immun. 2: 335-342. [Medline]
  58. Prossnitz, E. R.. 1997. Desensitization of N-formylpeptide receptor-mediated activation is dependent upon receptor phosphorylation. J. Biol. Chem. 272: 15213-15219. [Abstract/Free Full Text]
  59. Seligmann, B. E., M. P. Fletcher, J. I. Gallin. 1982. Adaptation of human neutrophil responsiveness to the chemoattractant N-formylmethionylleucylphenylalanine. J. Biol. Chem. 257: 6280-6286. [Abstract/Free Full Text]

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