Various inflammatory diseases are characterized by tissue infiltration of neutrophils. Chemokines recruit and activate leukocytes, but neutrophils are traditionally known to be restricted in their chemokine receptor (CR) expression repertoire. Neutrophils undergo phenotypic and functional changes under inflammatory conditions, but the mechanisms regulating CR expression of infiltrated neutrophils at sites of chronic inflammation are poorly defined. Here we show that infiltrated neutrophils from patients with chronic inflammatory lung diseases and rheumatoid arthritis highly express CR on their surface that are absent or only marginally expressed on circulating neutrophils, i.e., CCR1, CCR2, CCR3, CCR5, CXCR3, and CXCR4, as measured by flow cytometry, immunohistochemistry, and confocal microscopy. The induction of CR surface expression on infiltrated neutrophils was functionally relevant, because receptor activation by chemokine ligands ex vivo modulated neutrophil effector functions such as respiratory burst activity and bacterial killing. In vitro studies with isolated neutrophils demonstrated that the surface expression of CR was differentially induced in a cytokine-mediated, protein synthesis-dependent manner (CCR1, CCR3), through Toll-like (CXCR3) or NOD2 (CCR5) receptor engagement, through neutrophil apoptosis (CCR5, CXCR4), and/or via mobilization of intracellular CD63+ granules (CXCR3). CR activation on infiltrated neutrophils may represent a key mechanism by which the local inflammatory microenvironment fine-tunes neutrophil effector functions in situ. Since the up-regulation of CR was exclusively found on infiltrated neutrophils at inflammatory sites in situ, the targeting of these G protein-coupled receptors may have the potential to site-specifically target neutrophilic inflammation.

Neutrophils provide the first and most potent cellular line of innate host defense (1, 2). Leukocytes migrate to sites of inflammation via chemokines (3), which act through seven-transmembrane domain G protein-coupled receptors termed chemokine receptors. Seven CXC (CXCR1–7), 10 CCR (CCR1–10), 1 CX3CR (CX3CR1), and 1 CR (XCR1) chemokine receptor have been identified so far (4). Chemokines can be divided into inflammatory and homeostatic chemokines (5). Inflammatory chemokines are expressed in inflamed tissues and are critical for attracting effector leukocytes, whereas homeostatic chemokines maintain physiological traffic and immune surveillance by leukocytes (5).

In contrast to lymphocytes, monocytes and eosinophils (6, 7), human neutrophils are traditionally known to express only a very limited number of chemokine receptors (8, 9, 10). While CC chemokines mainly interact with lymphocytes, macrophages, and monocytes (11), neutrophils express predominantly receptors of the CXC or CX3C family, in particular CXCR1 and CXCR2 (12). Except for CXCR1/CXCR2 ligands, the majority of chemokines has been suggested to have no functional effect on human neutrophils (13). Evidence from animal models, however, suggests that under inflammatory conditions neutrophils undergo phenotypic changes driven by the surrounding microenvironment (1, 14), enabling them to expand their chemokine receptor expression pattern and respond to chemokines that are functionally inactive under resting conditions (15, 16, 17, 18). Studies with isolated human neutrophils have further demonstrated that GM-CSF, IFN-γ, or TNF-α are capable of priming neutrophils for migration to CC chemokines, mediated via up-regulation of CCR1 and CCR3 expression and/or via a CCR5-mediated mechanism (8, 19, 20, 21). However, the chemokine receptor expression and functionality of infiltrated neutrophils at sites of chronic inflammation and the underlying mechanisms regulating chemokine receptor expression on infiltrated neutrophils are poorly understood.

We investigated whether infiltrating neutrophils in human inflammatory diseases gain a new chemokine receptor expression profile and whether this has functional consequences. Therefore, we analyzed the expression and function of a broad variety of chemokine receptors on neutrophils directly isolated from sites of inflammation, i.e., from bronchoalveolar lavage fluid (BALF)2 of patients with chronic inflammatory lung diseases and from synovial fluid (SF) of patients with rheumatoid arthritis (RA). Our results indicate that infiltrated neutrophils display a novel chemokine receptor expression profile and migrate to the respective chemokines. Furthermore, chemokine activation of isolated infiltrated neutrophils resulted in a modulation of neutrophil effector functionality ex vivo. Several components of the inflammatory microenvironment, in particular cytokines, TLR ligands, NOD2 ligands, and apoptosis, were found to differentially induce the chemokine receptor expression on neutrophils via protein neosynthesis or granule mobilization. These studies suggest inducible chemokine receptors on neutrophils as functional and site-specific therapeutic targets in chronic inflammatory diseases.

Recombinant human IL-8, TNF-α, IL-1β, IFN-γ, and GM-CSF were from PeproTech. PMA, platelet-activating factor (PAF), fMLP, cytochalasin B, AMD3100, cycloheximide (CHX), LPS (from Escherichia coli serotype 0111:B4), lipoteichoic acid (LTA) from Staphylococcus aureus, and zymosan A (Saccharomyces cerevisiae) were from Sigma-Aldrich. Purified flagellin (Salmonella typhimurium) was from Alexis. Nonmethylated CpG motif-containing DNA (CpG) (GGTGCATCGATGCAGGGGGG) was from Invitrogen. R848 (resiquimod hydrochloride) was from GL Synthesis. Polyinosine-polycytidylic acid (poly(I:C); dsRNA) was from Pharmacia. Peptidoglycan (PGN; from S. aureus) was from InvivoGen. Muramyl dipeptide (MDP) was from Calbiochem. Pam3CysSerLys4 (Pam3CSK4) was from EMC Microcollections. Human serum albumin and fibronectin were obtained from Sanquin. RPMI 1640 was from Life Technologies. Recombinant human CCL2, CCL4, CCL11, CCL15, CXCL11, and CXCL12 were from R&D Systems. Mouse anti-human CCR1 and mouse anti-human CCR3 blocking Abs were from MBL International. A mouse anti-human CCR5-blocking Ab was from Sigma-Aldrich. A mouse anti-human CCR2-blocking Ab was from Abcam. A mouse anti-human CXCR3-blocking Ab was from Lifespan Biosciences. All reagents, buffers, and media were free of LPS (<0.01 ng/ml) by Limulus assay (Sigma-Aldrich).

The group of chronic lung diseases comprised patients with cystic fibrosis (CF), chronic obstructive pulmonary disease (COPD) and asthma (Table I). The CF group included five male and five female patients. Inclusion criteria for CF patients were the diagnosis of CF by clinical symptoms and positive sweat tests or disease-inducing mutations, forced expiratory volume in 1 s (FEV1) >25% of predicted value, and being on stable concomitant therapy at least 2 wk before the study. The CF patients had moderate to severe symptoms of the disease, as defined by the activity and physical examination criteria of the scoring system of Shwachman and Kulczycki (22). Six CF patients inhaled recombinant human DNase and four patients inhaled bronchodilators. None of the CF patients received inhaled or systemic corticosteroids or had signs of a severe systemic infection within 2 mo before the study. All CF patients were clinically stable at least 2 mo before the study, as indicated by lack of self-reported change in symptoms over the preceding 2 mo, and none reported a change in airway symptoms in the 2 mo before the study. The COPD group included four male and six female patients. Inclusion criteria were the clinical diagnosis of COPD, defined in accordance with American Thoracic Society criteria as a >2-year history of daily cough productive of phlegm for at least 3 mo of the year (23). All COPD patients suffered from chronic bronchitis with sputum production for at least 3 mo during 2 successive years. According to the Global Initiative for Chronic Obstructive Lung Disease stage classification, these patients had COPD stage II (24). All COPD patients were current smokers or had smoked previously. Five COPD patients used inhaled bronchodilators. All COPD patients were clinically stable at least 2 mo before the study, as indicated by lack of self-reported change in symptoms over the preceding 2 mo, and none reported a change in airway symptoms in the 2 mo before the study. The asthma group included five male and six female patients with allergic asthma. Inclusion criteria were recurrent episodes of wheezing and objective evidence of asthma as indicated by β2 agonist-reversible airflow obstruction (≥12% improvement in FEV1% predicted), bronchial hyperresponsiveness (exercise challenge), and ≥20% intraday peak flow variability, elevated total serum IgE (>150 kU/ml), and/or the presence of specific IgE (RAST class >2). The RAST was performed for 40 inhalation and food allergens. All asthma patients used inhaled bronchodilators. Spirometry and flow volume curves were performed in all patients according to the American Thoracic Society guidelines (25). Ten patients with RA (six males, four females) were included in the study. RA was diagnosed according to American College of Rheumatology criteria (26). All RA patients had at least one swollen knee joint and had active rheumatic disease, with a mean (±SD) serum C-reactive protein level of 39 ± 23 mg/L and an ESR of 51 ± 18 mm/hour. Ten age-matched healthy subjects were selected as the control group (six males, four females; mean age). These subjects were all nonsmokers, had no suspected or proven pulmonary disease, and were free of respiratory tract infections. The control subjects underwent minor surgical interventions and bronchoalveolar lavage was performed before the surgical procedure. The study was approved by the institutional review board of the Medical Faculty, Ludwig-Maximilians University (Munich, Germany). Written informed consent was obtained from all patients and healthy control subjects before enrollment.

Table I.

Patient characteristicsa

CFCOPDAsthmaRAControls
No. 10 10 11 10 10 
Age (years) 24 ± 5 62 ± 4 41 ± 8 60 ± 12 23 ± 8 
Sex (male/female) 5/5 4/6 5/6 6/4 6/4 
FEV1 (% predicted) 74 ± 18 71 ± 18 79 ± 14 ND ND 
Forced vital capacity (% predicted) 82 ± 17 81 ± 16 86 ± 12 ND ND 
No. using inhaled corticosteroids (μg/day) 3 (700 ± 180) 8 (1000 ± 290) 
Neutrophils (%) in BALF/SF 63 ± 13 27 ± 9 21 ± 11 74 ± 13 5 ± 2 
CFCOPDAsthmaRAControls
No. 10 10 11 10 10 
Age (years) 24 ± 5 62 ± 4 41 ± 8 60 ± 12 23 ± 8 
Sex (male/female) 5/5 4/6 5/6 6/4 6/4 
FEV1 (% predicted) 74 ± 18 71 ± 18 79 ± 14 ND ND 
Forced vital capacity (% predicted) 82 ± 17 81 ± 16 86 ± 12 ND ND 
No. using inhaled corticosteroids (μg/day) 3 (700 ± 180) 8 (1000 ± 290) 
Neutrophils (%) in BALF/SF 63 ± 13 27 ± 9 21 ± 11 74 ± 13 5 ± 2 
a

Results are expressed as means ± SD.

Bronchoscopy and bronchoalveolar lavage (4 × 1 ml of 0.9% (w/v) NaCl/kg body weight) were performed as described previously (27). The obtained BALF was filtered through two layers of sterile gauze. The first fraction of BALF was used because it contains higher percentages of neutrophils compared with the pooled fraction (28). The sample processing was performed immediately on ice. After centrifugation (200 × g, 10 min), the supernatant was stored at −80°C until analysis. The cell pellet was resuspended in 5 ml of PBS and used for preparation of cytospin slides and flow cytometry.

Paraffin-embedded, 5-μm-thick lung tissue sections of three COPD patients were stained. Slides were deparaffinized through a series of xylene baths and rehydrated through graded alcohols. The sections were then immersed in methanol containing 0.3% hydrogen peroxide for 20 min to block endogenous peroxidase activity and incubated in 2.5% blocking serum to reduce nonspecific binding. Sections were incubated overnight at 4°C with primary mouse anti-CCR1, anti-CCR2, anti-CCR3, anti-CCR5, anti-CXCR3, or anti-CXCR4 Abs (R&D Systems) at dilutions of 1/1000. The sections were then incubated with secondary anti-mouse Abs and underwent standard avidin-biotin immunohistochemical staining according to the manufacturer’s recommendations (Vector Laboratories). Diaminobenzidine was used as a chromogen and hematoxylin was used for counterstaining.

SF was obtained from patients with RA who presented with arthritis of the knee for diagnostic or therapeutic arthrocentesis. SF leukocytes were differentiated routinely by light microscopy after Jenner-Giemsa staining. RA-SF were centrifuged at 4°C (3 min; 2000 × g) and the cell-free supernatants were stored at −80°C until use.

The following mAbs were used: CD16-allophycocyanin mouse IgG1, CCR1-PE mouse IgG2b, CCR2-PE mouse IgG2b, CCR3-PE rat IgG2a, CCR4-PE IgG2b, CCR5-PE mouse IgG2b, CXCR3-FITC mouse IgG1 (all R&D Systems) and CD63-PE mouse IgG1, CD66b-PE mouse IgG2a, CD35-PE mouse IgG1, CD15-FITC mouse IgM, and CXCR4-PE mouse IgG2a (all from BD Pharmingen). Mouse IgG1-allophycocyanin, mouse IgG1-PE, rat IgG2a-PE, mouse IgM-FITC, mouse IgG1-CyChrome, mouse IgG2a-PE, and mouse IgG2b-PE (Immunotech) were used as isotype controls and were subtracted from the respective specific Ab staining. The results were expressed as mean fluorescence intensity (MFI). Calculations were performed with CellQuest analysis software (BD Biosciences).

Neutrophils from peripheral blood, BALF, or SF were preincubated for 30 min at 4°C with 20% (v/v) serum to prevent nonspecific binding via Fc receptors. Afterward, neutrophils were incubated with mAbs for 40 min, washed two times, and analyzed by flow cytometry as described previously (29). Neutrophils were identified by their light scatter properties and the expression of CD15 and CD16. Propidium iodide (PI, 5 μg/ml; Sigma-Aldrich) and Annexin VFITC (5 μg/ml; Boehringer Mannheim) were used to discriminate viable leukocytes (annexin VPI) from apoptotic (annexin V+PI) and necrotic (annexin V+PI+) leukocytes. These gates were used to analyze 10,000 neutrophils in blood, BALF, or SF from each sample. The gating of neutrophils vs eosinophils and macrophages was optimized before the study. For some experiments, neutrophils were permeabilized before being subjected to flow cytometry. Washed cells were fixed and permeabilized with the IntraPrep intracellular staining kit (Beckman Coulter) and incubated for 20 min according to the manufacturer’s instructions.

Neutrophils isolated from BALF of COPD patients were washed twice, resuspended in 100 μl of HBSS, and fixed in an equal volume of 4% (v/v) paraformaldehyde for 30 min. Cells were permeabilized with an equal volume of cold 0.1% (w/v) Triton X-100 for 2 min on ice. After washing and resuspending in 100 μl of HBSS, neutrophils were incubated with an optimal concentration of anti-CCR1 mouse IgG2b, anti-CCR2 mouse IgG2b, anti-CCR3 mouse IgG2b, anti-CCR5 mouse IgG2b, anti-CXCR3 mouse IgG1 and anti-CXCR4 IgG2a (R&D Systems) or the respective isotype controls. After staining with anti-mouse Alexa Fluor 555 along with Con A (lectin that binds to cell membranes) conjugated to Alexa Fluor 488, confocal laser-scanning microscopy was performed with the Leica TCS NT laser system, including a Leica DM IRB microscope with ×63 objectives. Cross-talk between the green and red channel was avoided by use of sequential scanning. Luminosity analysis was performed with SigmaScan Pro software.

Neutrophils were isolated from peripheral blood according to standard procedures (30). Venous blood was drawn and neutrophils were isolated immediately by density gradient centrifugation over isotonic Percoll according to the manufacturer’s instructions (Pharmacia). After lysis of the erythrocytes, the neutrophils were harvested, washed twice with HBSS (Sigma-Aldrich) containing 20 mM HEPES (Sigma- Aldrich) and 0.1% (w/v) BSA (Sigma-Aldrich), and resuspended in HBSS, 20 mM HEPES, and 0.1% (w/v) BSA at a cell concentration of 106/ml. The purity of the neutrophil suspensions was ≥98%, as assessed by microscopic examination of Pappenheim cytospin preparations. Cell viability was confirmed by trypan blue dye exclusion (≥95%).

Neutrophils from BALF and SF were isolated as described previously (31). The obtained BALF and SF cells were washed twice with sterile HBSS and were then resuspended in 10 ml of HBSS. These cells were layered onto a Percoll gradient (40 and 50% (w/v) solutions in saline) and centrifuged for 20 min at 400 × g to remove macrophages, epithelial cells, and lymphocytes. Then, the neutrophils were recovered from the interface of the 50 and 40% phases. The purity of the neutrophil suspensions was ≥98%, as assessed by May-Grünwald-Giemsa staining. Cell viability was confirmed by trypan blue dye exclusion (≥95%).

Levels of CCL2, CCL4, CCL11, CCL15, CXCL11, CXCL12, TNF-α, GM-CSF, and IFN-γ were determined in BALF and SF by a sandwich ELISA (R&D Systems) according to the manufacturer’s instructions.

Neutrophil chemotaxis was analyzed as described by a modified method of De Gendt et al. (32). In brief, isolated neutrophils were labeled with 4 μg/ml calcein-acetomethylester in HEPES medium for 45 min at 37°C before the start of the assay. After labeling, the cells were washed twice and resuspended in HBSS (106/ml). Calcein-labeled neutrophils (0.5 × 106 cells) were placed in the upper compartment and CCL2, CCL4, CCL11, CCL15, CXCL11, or CXCL12 (each 100 nM) were placed in the lower compartment of a Transwell filter system (3.0-μm pore size, 12-mm diameter; Costar). The Transwells were incubated for 60 min at 37°C. The percentage of cell migration was calculated by cell fluorescence, i.e., the amount of fluorescence in each of the compartments was measured and related to the fluorescence of the total input (set at 100%).

Chemokinesis was evaluated as described previously by Schweizer and coworkers (33). In brief, Transwell chambers were used to discriminate chemokinesis from chemotaxis. We added the chemokines CCL2, CCL3, CCL4, CCL11, CXCL11, or CXCL12 at different concentrations either in the lower, the upper, or both compartments of a Transwell chamber and analyzed the resulting migratory capacity of COPD BALF-isolated neutrophils. Cell migration was analyzed via checkerboard analysis. When migration was induced only with chemokines in the lower chamber, gradient-dependent migration (“chemotaxis”) was assumed. When cell migration was also observed with chemokines in the upper-only or upper plus lower chambers present, nondirected/gradient-independent migration (“chemokinesis”) was assumed.

Neutrophils were isolated from peripheral blood of healthy controls (2 × 106/ml). Thereupon, neutrophils were incubated for 6 h at 37°C with cell-free BALF supernatant, cell-free SF supernatant, GM-CSF (100 ng/ml), IL-8 (72 aa, 50 ng/ml), TNF-α (50 ng/ml), IL-1β (100 ng/ml), IFN-γ (1000 U/ml), or the combination of GM-CSF (10 ng/ml), TNF-α (100 ng/ml), and IFN-γ (1000 U/ml). The doses of these agents were chosen based on experiments before the study, showing maximal activation of human neutrophils without affecting cell viability. To examine whether the induction of chemokine receptor expression on neutrophils is dependent on protein synthesis, the effect of CHX (2 μg/ml) pretreatment was analyzed. After the incubation period, neutrophils were washed twice with ice-cold PBS, incubated with CCR1, CCR2, CCR3, CCR5, CXCR3, and CXCR4 Abs for 40 min (see above), washed two times with PBS, and analyzed by flow cytometry as described above.

α-Defensins were quantified in cell-free supernatants by sandwich ELISA with anti-human α-defensin mAb, anti-human α-defensin polyclonal Ab, HRP-conjugated rabbit anti-goat IgG, and recombinant human α-defensin-1 as described previously (34).

Neutrophils were isolated from peripheral blood and BALF of COPD patients and SF of RA patients. Chemokine receptors were blocked by pretreatment of neutrophils with anti-CCR1-, anti-CCR2-, anti-CCR3-, anti-CCR5-, anti-CXCR3-blocking Abs (each 10 μg/ml) or AMD3100 (3 μM) for 30 min. P. aeruginosa bacteria were incubated with the fluorescent ligand Lucifer Yellow (LY) for 60 min at room temperature. After preopsonization (60 min, 37°C, 20% pooled fresh C5a-depleted human serum), the bacteria (2 × 107/ml) in the HBSS-gel were incubated at 37°C for 2 h with neutrophils (2 × 106/ml) in the absence or presence of CCL2, CCL4, CCL11, CCL15, CXCL11, or CXCL12 (each 100 nM). Thereafter, the neutrophils were separated from the free bacteria by three centrifugations at 200 × g for 5 min. The LY fluorescence of the isolated neutrophils was analyzed by flow cytometry.

The respiratory burst of isolated neutrophils was analyzed by flow cytometry. Dihydrorhodamine 123 (DHR, Molecular Probes) was dissolved in DMSO at a concentration of 500 pM. After washing, aliquots (1 ml) of the isolated neutrophils (2 × 106/ml) were incubated with DHR for 20 min at 37°C. After several washings to remove unincorporated DHR, neutrophils were treated with CCL3, CCL2, CCL11, CCL5, CXCL11, or CXCL12 (each 100 nM) for 30 min at 37°C to activate the CCR1, CCR2, CCR3, CCR5, CXCR3, or CXCR4 function. After the chemokine stimulation, fMLP (1 μM) was added for 5 min at 37°C. Afterward, the reactions were stopped by transferring the sample tubes onto ice, and cells were immediately analyzed on the flow cytometer as described previously (35). Where indicated, neutrophils were pretreated for 30 min with anti-CCR1-, anti-CCR2-, anti-CCR3-, anti-CCR5-, anti-CXCR3-blocking mAbs (each 10 μg/ml) or AMD3100 (3 μM). Background MFI from a control tube without stimulation was subtracted from the MFI of the stimulated cells. Results are expressed as the MFI of the total neutrophil population. Each complete experiment was conducted three times.

For bacterial killing, a modified method according to Berger et al. (36) was used. A clinical isolate of a mucoid P. aeruginosa from a CF patient’s sputum was subcultured overnight, grown to stationary phase, washed, and preopsonized by incubation for 60 min at 37°C in 20% pooled, fresh C5a-depleted human serum. After washing two times in PBS, the opsonized P. aeruginosa bacteria were resuspended in 1 ml of a mixture of HBSS-gel and tryptic soy broth (Difco Laboratories). Afterward, neutrophils isolated from BALF of a patient with COPD were mixed with preopsonized bacteria (2 × 107 bacteria/ml) at a ratio of 1:5 (neutrophils/bacteria) in the absence or presence of CCL2, CCL4, CCL11, CCL15, CXCL11, or CXCL12 (each 100 nM). Aliquots of each mixture were removed immediately and after 30, 60, 90, and 120 min of incubation at 37°C. P. aeruginosa colonies at each time interval were counted by serial dilution in distilled water and quantitative spread plating and were expressed as CFU per ml. At 60 min, anti-CXCR3-blocking Abs (each 10 μg/ml) or AMD3100 (3 μM) were added to the CXCL12- or CXCL11-treated aliquots.

The effect of the TLR/NOD2 ligands LPS (100 ng/ml; TLR4), Pam3CSK4 (10 μg/ml; an artificial triacylated lipopeptide; TLR1/2), PGN (10 μg/ml; TLR2/NOD2), MDP (10 μg/ml; a component of PGN that binds to NOD2), LTA (10 μg/ml; TLR2), R-848 (10 μg/ml; a ssRNA analog; TLR7/8), poly(I:C) (50 μg/ml; a dsRNA analog; TLR3), CpG-DNA (100 μg/ml; TLR9), flagellin (1 μg/ml; TLR5), and zymosan (50 μg/ml; TLR2/6) was assessed on CCR1, CCR2, CCR3, CCR5, CXCR3, or CXCR4 expression on neutrophils. The doses of these agents were chosen based on experiments before the study showing maximal activation of human neutrophils without affecting cell viability. Since the response to TLR agonists has been found to be enhanced by GM-CSF pretreatment (37), neutrophils isolated from peripheral blood of healthy controls were preincubated with 50 ng/ml recombinant human GM-CSF for 90 min and then treated with the TLR ligands for 6 h at 37°C. After the incubation period, neutrophils were washed twice with ice-cold PBS, incubated with CCR1, CCR2, CCR3, CCR5, CXCR3, and CXCR4 Abs for 40 min (see above), washed two times with PBS, and analyzed by flow cytometry as described above.

Experiments were performed in duplicates and repeated at least three times. For parametric data, means ± SEM are given and the two-sided t test performed. A p < 0.05 was regarded as significant (SPSS statistical program, version 11.5) (38).

We gated infiltrated pulmonary and synovial neutrophils based on their light scatter (FSC/SCC) characteristics and based on positive expression of CD16 and CD15 (Fig. 1). This combined approach was able to differentiate neutrophils (FSClowCD15highCD16high) from alveolar macrophages (FSChighCD15lowCD16intermed) and infiltrated eosinophils (FSClowCD15intermedCD16low). We stained these neutrophils for a broad variety of CC and CXC chemokine receptors (CCR1–CCR5, CXCR1–4). To confirm the cellular identity and to avoid false-positive findings through contamination by non-neutrophils, we used immunohistochemistry and confocal microscopy (Fig. 2). These studies consistently demonstrated that infiltrated pulmonary and synovial neutrophils expressed CCR1, CCR2, CCR3, CCR5, CXCR3, and CXCR4, which were absent or marginally expressed on peripheral blood neutrophils (Figs. 2 and 3,A, upper panel). Neutrophils in BALF from healthy controls expressed low to intermediate levels of CCR1, CXCR3, and CXCR4 but expressed no detectable CCR2, CCR3, CCR4, or CCR5 (Fig. 3,A, upper panel). In contrast, BALF neutrophils from patients with chronic lung diseases as well as SF neutrophils from patients with RA had high surface expression levels of CCR1, CCR2, CCR3, CCR5, CXCR3, and CXCR4. These differences in chemokine receptor expression on infiltrated neutrophils were consistent when calculating either surface expression (MFI; Fig. 3,A, upper panel) or percentages of chemokine receptor-positive neutrophils (Fig. 3 A, lower panel). Patients with chronic lung diseases or RA had lower percentages of infiltrated CXCR1+ neutrophils compared with healthy controls or compared with peripheral blood. In BALF or SF, lower percentages of infiltrated CXCR2+ neutrophils were found compared with peripheral blood.

FIGURE 1.

Neutrophil gating. Neutrophils in BALF or SF were gated according to the following method (as shown here for BAL cells). First, granulocytes were gated based on their light scatter characteristics (FSC/SSC; upper panel). Gran, Granulocytes; AMs, alveolar macophages; Lyc, lymphocytes. The granulocyte region was further differentiated by means of the leukocyte surface markers CD16 and CD15 (lower panel). Combining the FSC/SSC and CD15/CD16 gating strategy, neutrophils (PMN) (FSClowCD15highCD16high) were differentiated from alveolar macrophages (FSChighCD15lowCD16intermed) and eosinophils (Eos) (FSClowCD15intermedCD16low).

FIGURE 1.

Neutrophil gating. Neutrophils in BALF or SF were gated according to the following method (as shown here for BAL cells). First, granulocytes were gated based on their light scatter characteristics (FSC/SSC; upper panel). Gran, Granulocytes; AMs, alveolar macophages; Lyc, lymphocytes. The granulocyte region was further differentiated by means of the leukocyte surface markers CD16 and CD15 (lower panel). Combining the FSC/SSC and CD15/CD16 gating strategy, neutrophils (PMN) (FSClowCD15highCD16high) were differentiated from alveolar macrophages (FSChighCD15lowCD16intermed) and eosinophils (Eos) (FSClowCD15intermedCD16low).

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FIGURE 2.

Chemokine receptor expression on pulmonary neutrophils. Chemokine receptor expression on pulmonary neutrophils was assessed by flow cytometry (left column), immunohistochemistry (middle column), and confocal microscopy (right column). Flow cytometry: representative histograms are shown of neutrophils from BALF of a representative patient with COPD and a representative control subject (Control) and from peripheral blood (Blood) of a control subject. Isotype expression overlapped with the specific expression in peripheral blood. Immunohistochemistry: representative sections of lung tissue from three COPD patients are shown. Diaminobenzidine was used as chromogen and hematoxylin for counterstaining. Black arrows mark chemokine receptor-expressing neutrophils. Confocal microscopy: Neutrophils isolated from BALF from COPD patients were fixed in paraformaldehyde and permeabilized with Triton X-100. After washing, the neutrophils were incubated with mouse anti-CXCR3, anti-CXCR4, anti-CCR1, anti-CCR2, anti-CCR3, or anti-CCR5 or the respective isotype controls. After staining with anti-mouse Alexa Fluor 555 along with Con A conjugated to Alexa Fluor 488, confocal laser-scanning microscopy was performed with ×63 objectives. Red color represents chemokine receptors and green color shows Con A (membrane and cytoplasm staining).

FIGURE 2.

Chemokine receptor expression on pulmonary neutrophils. Chemokine receptor expression on pulmonary neutrophils was assessed by flow cytometry (left column), immunohistochemistry (middle column), and confocal microscopy (right column). Flow cytometry: representative histograms are shown of neutrophils from BALF of a representative patient with COPD and a representative control subject (Control) and from peripheral blood (Blood) of a control subject. Isotype expression overlapped with the specific expression in peripheral blood. Immunohistochemistry: representative sections of lung tissue from three COPD patients are shown. Diaminobenzidine was used as chromogen and hematoxylin for counterstaining. Black arrows mark chemokine receptor-expressing neutrophils. Confocal microscopy: Neutrophils isolated from BALF from COPD patients were fixed in paraformaldehyde and permeabilized with Triton X-100. After washing, the neutrophils were incubated with mouse anti-CXCR3, anti-CXCR4, anti-CCR1, anti-CCR2, anti-CCR3, or anti-CCR5 or the respective isotype controls. After staining with anti-mouse Alexa Fluor 555 along with Con A conjugated to Alexa Fluor 488, confocal laser-scanning microscopy was performed with ×63 objectives. Red color represents chemokine receptors and green color shows Con A (membrane and cytoplasm staining).

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FIGURE 3.

A, Neutrophil chemokine receptor expression in peripheral blood, BALF, and SF. Chemokine receptor expression levels on neutrophils from peripheral blood of healthy controls (□) or patients with lung diseases (white hatched bars), from BALF of healthy controls (▦) or patients with lung diseases (gray hatched bars) and from peripheral blood and SF of patients with RA (▪). Receptor expression on neutrophils was analyzed by flow cytometry. The upper panel shows the surface expression levels of chemokine receptors, the lower panel shows percentages of chemokine receptor-expressing neutrophils. Bars represent means ± SEM. ∗, p < 0.05: vs neutrophils in peripheral blood. B, Chemokines in bronchoalveolar lavage and synovial fluid. Levels of human CCL2, CCL3, CCL4, CCL5, CCL11, CXCL11, and CXCL12 were quantified in BALF of healthy controls, in BALF of patients with lung diseases, and in SF of patients with RA by sandwich ELISA. Bars represent means ± SEM. ∗, p < 0.05 vs BALF from healthy controls. C, Neutrophil chemokine receptor expression in BALF from patients with chronic inflammatory lung diseases. Chemokine receptor expression levels on neutrophils in BALF of healthy controls and patients with CF, COPD, or allergic asthma. Receptor expression on neutrophils was analyzed by flow cytometry. Bars represent means ± SEM. ∗, p < 0.05 vs neutrophils in peripheral blood.

FIGURE 3.

A, Neutrophil chemokine receptor expression in peripheral blood, BALF, and SF. Chemokine receptor expression levels on neutrophils from peripheral blood of healthy controls (□) or patients with lung diseases (white hatched bars), from BALF of healthy controls (▦) or patients with lung diseases (gray hatched bars) and from peripheral blood and SF of patients with RA (▪). Receptor expression on neutrophils was analyzed by flow cytometry. The upper panel shows the surface expression levels of chemokine receptors, the lower panel shows percentages of chemokine receptor-expressing neutrophils. Bars represent means ± SEM. ∗, p < 0.05: vs neutrophils in peripheral blood. B, Chemokines in bronchoalveolar lavage and synovial fluid. Levels of human CCL2, CCL3, CCL4, CCL5, CCL11, CXCL11, and CXCL12 were quantified in BALF of healthy controls, in BALF of patients with lung diseases, and in SF of patients with RA by sandwich ELISA. Bars represent means ± SEM. ∗, p < 0.05 vs BALF from healthy controls. C, Neutrophil chemokine receptor expression in BALF from patients with chronic inflammatory lung diseases. Chemokine receptor expression levels on neutrophils in BALF of healthy controls and patients with CF, COPD, or allergic asthma. Receptor expression on neutrophils was analyzed by flow cytometry. Bars represent means ± SEM. ∗, p < 0.05 vs neutrophils in peripheral blood.

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These increased receptor expressions on BALF and SF neutrophils were paralleled by increased levels of the corresponding chemokine ligands CCL2, CCL3, CCL4, CCL11, CCL15, CXCL11, and CXCL12 in BALF and SF (Fig. 3,B). CCR4 was not expressed on blood, BALF, or SF neutrophils. When the expression levels on BALF neutrophils were compared among patients with CF, COPD, and allergic asthma, asthmatics had the highest CCR3 and the lowest CCR5 and CXCR3 expression levels (Fig. 3 C). CF patients had lower CXCR4 expression levels and the highest CXCR3 expression compared with the other patient groups. Comparing patients with and without inhaled corticosteroids, we did not find statistically significant differences (data not shown). These results indicate that 1) infiltrated neutrophils express novel chemokine receptors at sites of inflammation that are not present on peripheral blood neutrophils, 2) the induction of chemokine receptor expression is accompanied by increased levels of their corresponding chemokines, and 3) that different inflammatory diseases show distinct chemokine receptor expression patterns.

To examine whether the chemokine receptors expressed on infiltrated neutrophils in BALF and SF were functional, migration of BALF/SF-isolated neutrophils upon stimulation with their respective chemokine ligands was analyzed ex vivo (Fig. 4). Neutrophils isolated from BALF and SF migrated to their respective ligands, whereas neutrophils isolated from peripheral blood did not or only at a very low level. Ab blocking of the respective chemokine receptors largely abrogated the chemokine-induced neutrophil chemotaxis (Fig. 4,A), whereas isotype control Abs had no effect on chemokine-induced cell migration (data not shown). Since the migratory capacity of the ex vivo-isolated cells was relatively low compared with the capacity of freshly isolated neutrophils toward CXCR1/CXCR2 chemokines, we investigated whether the observed “chemotactic” effect was indeed gradient-dependent migration (chemotaxis) or could in fact be nondirected, gradient-independent migration (chemokinesis). Therefore, we performed studies to discriminate both mechanisms using a similar method as described previously (33). For this purpose, we added the chemokines CCL2, CCL3, CCL4, CCL11, CXCL11, or CXCL12 at different concentrations either in the lower, the upper, or in both compartments of a Transwell chamber and analyzed the resulting migratory capacity of COPD BALF-isolated neutrophils (Fig. 4 B). The migratory responses of all chemokines were dose-dependently induced with chemokines in the lower compartment of the chamber. When this chemoattractant was present in both compartments or just in the upper compartment of the chamber, no significant migratory responses were observed anymore, suggesting gradient-dependent chemotaxis.

FIGURE 4.

Chemokine receptors on infiltrated neutrophils are functional. A, Chemotaxis. Neutrophils were isolated from peripheral blood and BALF of COPD patients and SF of RA patients. Chemokine receptors were blocked by pretreatment of neutrophils with anti-CCR1-, anti-CCR2-, anti-CCR3-, anti-CCR5-, anti-CXCR3-blocking Abs (each 10 μg/ml) or AMD3100 (3 μM) for 30 min. Neutrophils (0.5 × 106 cells) were labeled with calcein and were placed in the upper compartment of a Transwell filter system. CCL2, CCL3, CCL4, CCL11, CXCL11, or CXCL12 (each at 100 nM) were placed in the lower compartment. The Transwells were incubated for 60 min at 37°C. The percent cell migration was calculated by flow cytometry, i.e., the amount of fluorescence in each of these compartments was measured and related to the fluorescence of the total input (set at 100%). Bars represent the means ± SEM of independent experiments with cells from three different donors. ∗, p < 0.05 Ab-treated vs nontreated neutrophils. B, Chemokinesis. B shows dose-dependent migratory responses of BALF-isolated neutrophils from COPD patients (n = 3). Chemotaxis was analyzed with the respective chemokines (CCL2, CCL3, CCL4, CCL11, CXCL11, or CXCL12 at the indicated concentrations) present in the lower compartment of the Transwell chamber (lower). Chemokinesis was analyzed with chemokines present in both compartments (both) or just in the upper compartment (upper) of the chamber. ∗, p < 0.05 lower chamber vs upper chamber or both chambers.

FIGURE 4.

Chemokine receptors on infiltrated neutrophils are functional. A, Chemotaxis. Neutrophils were isolated from peripheral blood and BALF of COPD patients and SF of RA patients. Chemokine receptors were blocked by pretreatment of neutrophils with anti-CCR1-, anti-CCR2-, anti-CCR3-, anti-CCR5-, anti-CXCR3-blocking Abs (each 10 μg/ml) or AMD3100 (3 μM) for 30 min. Neutrophils (0.5 × 106 cells) were labeled with calcein and were placed in the upper compartment of a Transwell filter system. CCL2, CCL3, CCL4, CCL11, CXCL11, or CXCL12 (each at 100 nM) were placed in the lower compartment. The Transwells were incubated for 60 min at 37°C. The percent cell migration was calculated by flow cytometry, i.e., the amount of fluorescence in each of these compartments was measured and related to the fluorescence of the total input (set at 100%). Bars represent the means ± SEM of independent experiments with cells from three different donors. ∗, p < 0.05 Ab-treated vs nontreated neutrophils. B, Chemokinesis. B shows dose-dependent migratory responses of BALF-isolated neutrophils from COPD patients (n = 3). Chemotaxis was analyzed with the respective chemokines (CCL2, CCL3, CCL4, CCL11, CXCL11, or CXCL12 at the indicated concentrations) present in the lower compartment of the Transwell chamber (lower). Chemokinesis was analyzed with chemokines present in both compartments (both) or just in the upper compartment (upper) of the chamber. ∗, p < 0.05 lower chamber vs upper chamber or both chambers.

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To examine whether activation of the chemokine receptors on neutrophils from BALF and SF also modulated antibacterial effector functions by neutrophils, respiratory burst, phagocytosis, and bacterial killing capacity of BALF/SF isolated neutrophils upon stimulation with their respective chemokines were analyzed. Although CCL2, CCL3, and CXCL11 treatment enhanced the respiratory burst of COPD BALF-isolated neutrophils toward fMLP stimulation, CXCL12 decreased the respiratory burst capacity (Fig. 5,A). Incubation with chemokines along with PMA gave similar results (data not shown). Similarly, CCL2, CCL3, and CXCL11 treatment increased the phagocytosis of P. aeruginosa (Fig. 5,B) by neutrophils, whereas CCL2, CCL3, CCL11, and CXCL11 stimulated the α-defensin release by neutrophils (Fig. 5,C). To investigate how these changes finally modulate the bacterial killing capacity of neutrophils, we incubated BALF-isolated neutrophils with these chemokines (Fig. 6). CXCL11 induced the strongest bacterial killing capacity, while CXCL12 attenuated bacterial killing (Fig. 6,A). Blocking of CXCR3 or CXCR4 reversed these effects (Fig. 6 B). These data indicate that chemokine receptor-expressing neutrophils in BALF are activated by their ligands, which results in an enhanced antibacterial capacity.

FIGURE 5.

Neutrophil effector functions. A, Respiratory burst. Neutrophils were isolated from peripheral blood and BALF of COPD patients and SF of RA patients. Chemokine receptors were blocked by pretreatment of neutrophils with chemokine receptor-blocking Abs (each 10 μg/ml) for 30 min. Neutrophils were loaded with DHR and were treated with CCL3, CCL2, CCL11, CCL5, CXCL11, or CXCL12 (each 100 nM) for 30 min at 37°C to activate the CCR1, CCR2, CCR3, CCR5, CXCR3, or CXCR4 function. After the chemokine stimulation, fMLP (1 μM) was added for 5 min at 37°C. Afterward, the reactions were stopped by transferring the sample tubes onto ice, and cells were immediately analyzed on the flow cytometer as described previously (35 ). Background MFI from a control tube without fMLP was subtracted from the MFI of the fMLP-stimulated cells. Results are expressed as the MFI of the total neutrophil population. ∗, p < 0.05 vs neutrophils from peripheral blood. B, Phagocytosis. Neutrophils were isolated from peripheral blood and BALF of COPD patients and SF of RA patients. Chemokine receptors were blocked by pretreatment of neutrophils with chemokine receptor-blocking Abs (each 10 μg/ml) for 30 min. P. aeruginosa bacteria were incubated with the fluorescent ligand LY for 60 min at room temperature. After preopsonization, the bacteria (2 × 107/ml) in HBSS-gel were incubated at 37°C for 2 h with neutrophils (2 × 106/ml) in the presence of the CCL2, CCL3, CCL4, CCL11, CXCL11, or CXCL12 (each 100 nM). After the incubation period, the neutrophils were separated from the free bacteria by three centrifugations at 200 × g for 5 min. The LY fluorescence of the isolated neutrophils was analyzed by flow cytometry. ∗, p < 0.05 vs neutrophils from peripheral blood. C, α-Defensin release. Neutrophils were isolated from peripheral blood and BALF of COPD patients and SF of RA patients. Chemokine receptors were blocked by pretreatment of neutrophils with chemokine receptor-blocking Abs (each 10 μg/ml) or AMD3100 (3 μM) for 30 min. Afterward, neutrophils were incubated with CCL2, CCL3, CCL4, CCL11, CXCL11, or CXCL12 (each at 100 nM) for 1 h. α-defensins were quantified in cell-free supernatants by sandwich ELISA with anti-human α-defensin mAb, anti-human α-defensin polyclonal Ab, HRP-conjugated rabbit anti-goat IgG, and recombinant human α-defensin-1. ∗, p < 0.05 vs neutrophils from peripheral blood. Treating neutrophils with isotype control Abs instead of blocking Abs had no effect on respiratory burst, phagocytosis, or α-defensin release (data not shown).

FIGURE 5.

Neutrophil effector functions. A, Respiratory burst. Neutrophils were isolated from peripheral blood and BALF of COPD patients and SF of RA patients. Chemokine receptors were blocked by pretreatment of neutrophils with chemokine receptor-blocking Abs (each 10 μg/ml) for 30 min. Neutrophils were loaded with DHR and were treated with CCL3, CCL2, CCL11, CCL5, CXCL11, or CXCL12 (each 100 nM) for 30 min at 37°C to activate the CCR1, CCR2, CCR3, CCR5, CXCR3, or CXCR4 function. After the chemokine stimulation, fMLP (1 μM) was added for 5 min at 37°C. Afterward, the reactions were stopped by transferring the sample tubes onto ice, and cells were immediately analyzed on the flow cytometer as described previously (35 ). Background MFI from a control tube without fMLP was subtracted from the MFI of the fMLP-stimulated cells. Results are expressed as the MFI of the total neutrophil population. ∗, p < 0.05 vs neutrophils from peripheral blood. B, Phagocytosis. Neutrophils were isolated from peripheral blood and BALF of COPD patients and SF of RA patients. Chemokine receptors were blocked by pretreatment of neutrophils with chemokine receptor-blocking Abs (each 10 μg/ml) for 30 min. P. aeruginosa bacteria were incubated with the fluorescent ligand LY for 60 min at room temperature. After preopsonization, the bacteria (2 × 107/ml) in HBSS-gel were incubated at 37°C for 2 h with neutrophils (2 × 106/ml) in the presence of the CCL2, CCL3, CCL4, CCL11, CXCL11, or CXCL12 (each 100 nM). After the incubation period, the neutrophils were separated from the free bacteria by three centrifugations at 200 × g for 5 min. The LY fluorescence of the isolated neutrophils was analyzed by flow cytometry. ∗, p < 0.05 vs neutrophils from peripheral blood. C, α-Defensin release. Neutrophils were isolated from peripheral blood and BALF of COPD patients and SF of RA patients. Chemokine receptors were blocked by pretreatment of neutrophils with chemokine receptor-blocking Abs (each 10 μg/ml) or AMD3100 (3 μM) for 30 min. Afterward, neutrophils were incubated with CCL2, CCL3, CCL4, CCL11, CXCL11, or CXCL12 (each at 100 nM) for 1 h. α-defensins were quantified in cell-free supernatants by sandwich ELISA with anti-human α-defensin mAb, anti-human α-defensin polyclonal Ab, HRP-conjugated rabbit anti-goat IgG, and recombinant human α-defensin-1. ∗, p < 0.05 vs neutrophils from peripheral blood. Treating neutrophils with isotype control Abs instead of blocking Abs had no effect on respiratory burst, phagocytosis, or α-defensin release (data not shown).

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FIGURE 6.

Bacterial killing. A, A clinical isolate of a mucoid P. aeruginosa from a CF patient’s sputum was subcultured overnight, grown to stationary phase, washed, and preopsonized by incubation for 60 min at 37°C in 20% pooled, fresh C5a-depleted human serum. After washing two times in PBS, the opsonized P. aeruginosa bacteria were resuspended in 1 ml of a mixture of HBSS-gel and tryptic soy broth. Afterward, neutrophils isolated from BALF of a patient with COPD were mixed with preopsonized bacteria (2 × 107 bacteria/ml) at a ratio of 1:5 (neutrophils/bacteria) in the absence or presence of CCL2, CCL4, CCL11, CCL15, CXCL11, or CXCL12 (each 100 nM). Aliquots of each mixture were removed after 0, 30, 60, 90, and 120 min of incubation at 37°C. P. aeruginosa colonies at each time interval were counted by serial dilution in distilled water and quantitative spread plating and were expressed as CFU per ml. B, After 60 min, anti-CXCR3-blocking Abs (10 μg/ml) or AMD3100 (3 μM) were added to the CXCL11- or CXCL12-treated aliquots. Treating neutrophils with isotype control Abs instead of blocking Abs had no effect on bacterial killing (data not shown). Bars represent the means ± SEM of three independent experiments with cells from different donors.

FIGURE 6.

Bacterial killing. A, A clinical isolate of a mucoid P. aeruginosa from a CF patient’s sputum was subcultured overnight, grown to stationary phase, washed, and preopsonized by incubation for 60 min at 37°C in 20% pooled, fresh C5a-depleted human serum. After washing two times in PBS, the opsonized P. aeruginosa bacteria were resuspended in 1 ml of a mixture of HBSS-gel and tryptic soy broth. Afterward, neutrophils isolated from BALF of a patient with COPD were mixed with preopsonized bacteria (2 × 107 bacteria/ml) at a ratio of 1:5 (neutrophils/bacteria) in the absence or presence of CCL2, CCL4, CCL11, CCL15, CXCL11, or CXCL12 (each 100 nM). Aliquots of each mixture were removed after 0, 30, 60, 90, and 120 min of incubation at 37°C. P. aeruginosa colonies at each time interval were counted by serial dilution in distilled water and quantitative spread plating and were expressed as CFU per ml. B, After 60 min, anti-CXCR3-blocking Abs (10 μg/ml) or AMD3100 (3 μM) were added to the CXCL11- or CXCL12-treated aliquots. Treating neutrophils with isotype control Abs instead of blocking Abs had no effect on bacterial killing (data not shown). Bars represent the means ± SEM of three independent experiments with cells from different donors.

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To examine which factors account for the induction of chemokine receptor surface expression on infiltrating neutrophils, we performed three approaches to mimic the inflammatory microenvironment infiltrated neutrophils are faced with: 1) proinflammatory cytokines, 2) long-term neutrophil culture/apoptosis, and 3) TLR/NOD2 activation.

Peripheral blood isolated neutrophils were incubated for 6 h with pooled cell-free BALF supernatant from CF patients, pooled cell-free SF supernatant from RA patients. or with proinflammatory cytokines that are typically found in BALF/SF of patients with chronic lung diseases or RA (Fig. 7,A). Both BALF and SF induced significant up-regulation of CXCR3 > CCR1 > CCR3 surface expressions on neutrophils. Further analyses revealed that GM-CSF and mainly IFN-γ-induced CCR1 expression, IFN-γ-induced CCR3 expression, and the combination of GM-CSF, TNF-α, and IFN-γ were the most potent stimulators of CCR1 and CCR3 surface expression. In contrast to CCR1 and CCR3, CXCR4 surface expression was decreased after treatment with these proinflammatory cytokines. Neither cytokine alone nor cytokines in combination had a significant effect on CCR2, CCR5, or CXCR3 surface expression on neutrophils. To evaluate whether the BALF (CXCR3)- or cytokine (CCR1, CCR3)-elicited up-regulation or down-regulation (CXCR4) of chemokine receptor expressions were dependent on protein neosynthesis, we used CHX. CHX pretreatment almost completely prevented the up-regulation of CCR1 and CCR3 expressions but had no effect on CXCR3 up- or CXCR4 down-regulation, suggesting differential involvement of protein neosynthesis. These studies indicate that proinflammatory cytokines differentially modulate chemokine receptor expression on neutrophils. Although CCR1 and CCR3 expressions are induced, CXCR4 expression is decreased upon cytokine stimulation. The stimulatory effect of BALF/SF on CXCR3 surface expression remained elusive. These data demonstrate that proinflammatory cytokines, in particular GM-CSF, TNF-α, and IFN-γ, differentially induce chemokine receptor expression on neutrophils. Therefore, we quantitated these cytokines in BALFs and SFs from the included patients and control subjects and found markedly increased levels of GM-CSF, TNF-α, and IFN-γ in all chronic disease conditions (Table II).

FIGURE 7.

Induction of chemokine receptor expression on neutrophils in vitro. A, Effect of BALF, SF, and proinflammatory cytokines on chemokine receptor expression of neutrophils. Neutrophils were isolated from peripheral blood of healthy controls (2 × 106/ml). Thereupon, neutrophils were incubated for 6 h at 37°C with cell-free BALF supernatant, cell-free SF supernatant, GM-CSF (10 ng/ml), IL-8 (500 ng/ml), TNF-α (100 ng/ml), IL-1β (100 ng/ml), IFN-γ (500 U/ml), or the combination of GM-CSF (10 ng/ml), TNF-α (100 ng/ml), and IFN-γ (500 U/ml). To examine whether the induction of chemokine receptor expression on neutrophils is dependent on protein synthesis, the effect of CHX (2 μg/ml) pretreatment was analyzed. After the incubation period, neutrophils were washed twice with ice-cold PBS, incubated with Abs, and analyzed by flow cytometry as described above. Bars represent the means ± SEM of independent experiments with cells from three different healthy control donors. ∗, p < 0.05 vs buffer-treated neutrophils. B, Effect of long-term incubation and cytokine stimulation on chemokine receptor expression of neutrophils. Neutrophils were isolated from peripheral blood of healthy controls (2 × 106/ml) and were incubated in serum-free HBSS for 0, 12, or 24 h at 37°C with or without CHX (2 μg/ml) pretreatment. After the incubation period, neutrophils were washed twice with ice-cold PBS, incubated with Abs for 40 min (see above), washed two times with PBS, and analyzed by flow cytometry as described above. In parallel, annexin V staining was performed. Bars represent the means ± SEM of independent experiments with cells from three different donors. ∗, p < 0.05 vs buffer-treated neutrophils. C, Annexin V. Peripheral blood neutrophils were in vitro cultured for 24 h. After the incubation period, neutrophils were incubated with Abs against CCR5 or CXCR4 along with annexin V (upper panel). BALF neutrophils were stained with CCR5, CXCR4, and/or annexin V directly after isolation (lower panel).

FIGURE 7.

Induction of chemokine receptor expression on neutrophils in vitro. A, Effect of BALF, SF, and proinflammatory cytokines on chemokine receptor expression of neutrophils. Neutrophils were isolated from peripheral blood of healthy controls (2 × 106/ml). Thereupon, neutrophils were incubated for 6 h at 37°C with cell-free BALF supernatant, cell-free SF supernatant, GM-CSF (10 ng/ml), IL-8 (500 ng/ml), TNF-α (100 ng/ml), IL-1β (100 ng/ml), IFN-γ (500 U/ml), or the combination of GM-CSF (10 ng/ml), TNF-α (100 ng/ml), and IFN-γ (500 U/ml). To examine whether the induction of chemokine receptor expression on neutrophils is dependent on protein synthesis, the effect of CHX (2 μg/ml) pretreatment was analyzed. After the incubation period, neutrophils were washed twice with ice-cold PBS, incubated with Abs, and analyzed by flow cytometry as described above. Bars represent the means ± SEM of independent experiments with cells from three different healthy control donors. ∗, p < 0.05 vs buffer-treated neutrophils. B, Effect of long-term incubation and cytokine stimulation on chemokine receptor expression of neutrophils. Neutrophils were isolated from peripheral blood of healthy controls (2 × 106/ml) and were incubated in serum-free HBSS for 0, 12, or 24 h at 37°C with or without CHX (2 μg/ml) pretreatment. After the incubation period, neutrophils were washed twice with ice-cold PBS, incubated with Abs for 40 min (see above), washed two times with PBS, and analyzed by flow cytometry as described above. In parallel, annexin V staining was performed. Bars represent the means ± SEM of independent experiments with cells from three different donors. ∗, p < 0.05 vs buffer-treated neutrophils. C, Annexin V. Peripheral blood neutrophils were in vitro cultured for 24 h. After the incubation period, neutrophils were incubated with Abs against CCR5 or CXCR4 along with annexin V (upper panel). BALF neutrophils were stained with CCR5, CXCR4, and/or annexin V directly after isolation (lower panel).

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Table II.

Cytokine levelsa

CF (n = 10)COPD (n = 10)Asthma (n = 11)RA (n = 10)Controls (n = 10)
TNF-α (pg/ml) 493 ± 272 312 ± 143 185 ± 77 265 ± 118 Not detectable 
GM-CSF (pg/ml) 547 ± 302 390 ± 187 273 ± 146 322 ± 226 29 ± 14 
IFN-γ (pg/ml) 54 ± 25 43 ± 39 18 ± 17 39 ± 28 Not detectable 
CF (n = 10)COPD (n = 10)Asthma (n = 11)RA (n = 10)Controls (n = 10)
TNF-α (pg/ml) 493 ± 272 312 ± 143 185 ± 77 265 ± 118 Not detectable 
GM-CSF (pg/ml) 547 ± 302 390 ± 187 273 ± 146 322 ± 226 29 ± 14 
IFN-γ (pg/ml) 54 ± 25 43 ± 39 18 ± 17 39 ± 28 Not detectable 
a

Results are expressed as means ± SD.

Since inflammatory neutrophils are long-lived compared with circulating neutrophils (1, 14, 39), we hypothesized that apoptosis/senescence modulates chemokine receptor expression on neutrophils. Our experiments showed that long-term culture of peripheral blood-isolated neutrophils induced CCR5 and CXCR4 surface expression after 12 h and most pronounced after 24 h of incubation (Fig. 7,B). Up-regulation of CCR5 and CXCR4 expression was largely prevented by CHX pretreatment. To test whether the induction of CCR5 and CXCR4 expression might be due to apoptosis, we assessed annexin V expression on cultured neutrophils (Fig. 7, B and C). After 12 h and more strongly after 24 h, the cultured neutrophils underwent apoptosis (Fig. 7,B). Although the majority of CCR5+ neutrophils were annexin V+, CXCR4+ neutrophils consisted of two almost equal populations of annexin V+ and annexin V cells (Fig. 7 C), suggesting that CXCR4 surface induction cannot be solely explained by apoptotic membrane changes. The induction of CCR5 and CXCR4 expression on the surface of neutrophils was completely (CCR5) or partially (CXCR4) inhibited by zVAD-fmk, a general apoptosis inhibitor (data not shown), suggesting a direct involvement of apoptosis in the up-regulation of CCR5 on neutrophils. In contrast to 24-h cultured peripheral blood-derived neutrophils, CCR5+ and CXCR4+ neutrophils in BALF from patients with COPD were mostly nonapoptotic (annexin V), indicating that factors in BALF might protect the neutrophils from apoptosis. When viewed in combination, these studies demonstrate that long-term incubation induces CCR5 and CXCR4 surface expression on neutrophils, effects that were dependent (CCR5) or partially dependent (CXCR4) on apoptosis.

Next, we investigated whether TLR or NOD2 ligands, which are abundantly present in CF and COPD airway fluids (40, 41), modulate chemokine receptor expression on isolated neutrophils. We treated neutrophils with TLR agonists plus GM-CSF, since GM-CSF was previously found to generally enhance the effect of TLR ligands on neutrophils (37). We found that among all chemokine receptors analyzed, only the surface expression of CCR5 and CXCR3 were significantly affected by treatment with TLR ligands (Fig. 8). The TLR2/NOD2 ligands PGN and MDP induced CCR5 expression, while the TLR1/TLR2 ligand Pam3CSK4 up-regulated CXCR3 surface expression on neutrophils. The up-regulation of CCR5 or CXCR3 surface expression after NOD2/TLR stimulation was not due to apoptosis or cell death as assessed by annexin V and PI staining (data not shown). GM-CSF slightly enhanced the effect of PGN, MDP, or Pam3CSK4 on CCR5 and CXCR3 expression, respectively. GM-CSF up-regulated CCR1 expression on neutrophils, as also shown above, but TLR ligands alone had no effect on CCR1 expression.

FIGURE 8.

Effect of Toll-like/NOD2 receptor ligands on chemokine receptor expression of neutrophils. Neutrophils (106/ml) from the blood of healthy controls were preincubated with 50 ng/ml recombinant human GM-CSF for 90 min and were then incubated for 6 h at 37°C in HBSS with 100 ng/ml LPS, 10 μg/ml Pam3CSK4, 10 μg/ml LTA, 10 μg/ml PGN, 10 μg/ml MDP, 10 μg/ml R-848, 50 μg/ml poly(I:C), 100 μg/ml CpG, 1 μg/ml flagellin, or 50 μg/ml zymosan A. After the incubation period, neutrophils were incubated with CCR1, CCR2, CCR3, CCR5, CXCR3, or CXCR4 Abs and analyzed by flow cytometry. The means ± SEM of five independent experiments with cells from different donors are shown. ∗, p < 0.05 vs HBSS-treated neutrophils.

FIGURE 8.

Effect of Toll-like/NOD2 receptor ligands on chemokine receptor expression of neutrophils. Neutrophils (106/ml) from the blood of healthy controls were preincubated with 50 ng/ml recombinant human GM-CSF for 90 min and were then incubated for 6 h at 37°C in HBSS with 100 ng/ml LPS, 10 μg/ml Pam3CSK4, 10 μg/ml LTA, 10 μg/ml PGN, 10 μg/ml MDP, 10 μg/ml R-848, 50 μg/ml poly(I:C), 100 μg/ml CpG, 1 μg/ml flagellin, or 50 μg/ml zymosan A. After the incubation period, neutrophils were incubated with CCR1, CCR2, CCR3, CCR5, CXCR3, or CXCR4 Abs and analyzed by flow cytometry. The means ± SEM of five independent experiments with cells from different donors are shown. ∗, p < 0.05 vs HBSS-treated neutrophils.

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No effects were found of apoptotic bystander cells (apoptotic T cells, neutrophils, or macrophages) on chemokine receptor expression of isolated neutrophils in vitro (data not shown).

When viewed in combination, these studies demonstrate that conditions typically present in the inflammatory microenvironment (cytokines, long-term culture and TLR ligands) differentially modulate the chemokine receptor expression on neutrophils.

Since the Pam3CSK4-induced up-regulation of CXCR3 surface expression occurred already after 30–60 min of TLR stimulation and was not prevented by CHX (data not shown), we hypothesized that CXCR3 might be stored intracellularly in neutrophils and does not require de novo receptor synthesis. Confocal microscopy of permeabilized neutrophils revealed that CXCR3 was stored intracellularly in resting neutrophils in a granular staining pattern (Fig. 9,A). The role of the microfilament and cytoskeletal apparatus in this process was evaluated with CytB, an inhibitor of microfilament functions, as described previously by us (42). Flow cytometric studies showed that stimulation of granule release with CytB plus fMLP, releasing primary and specific granules as well as secretory vesicles, strongly up-regulated CXCR3 surface expression on neutrophils, whereas PAF plus fMLP, releasing only specific granules and secretory vesicles, did not induce CXCR3 surface expression (Fig. 9,B). Further costainings with CD63, a marker for azurophil granules, CD66b, a marker for specific granules, or CD35, a marker for secretory vesicles, demonstrated coexpression of CXCR3 with CD63, but not with CD66b (Fig. 9 C) or with CD35 (data not shown). Stimulation with Pam3CSK4 also resulted in release of CD63+ granules associated with CXCR3 up-regulation on the cell surface (data not shown). These findings indicate that CXCR3 is stored in azurophil granules in neutrophils.

FIGURE 9.

CXCR3 expression in neutrophils. A, Confocal microscopy. Neutrophils were isolated from peripheral blood of healthy controls (2 × 106/ml). Where indicated, neutrophils were permeabilized with an equal volume of cold 0.1% (w/v) Triton X-100. Neutrophils were stained with rabbit anti-CXCR3 Abs, anti-rabbit-Ig Alexa Fluor 555, and Con A conjugated to Alexa Fluor 488. Confocal laser-scanning microscopy was performed with the Leica TCS NT laser system using a Leica DM IRB microscope. Red color represents CXCR3 receptors and green color Con A. B, Mobilization of intracellular CXCR3 expression. Neutrophils (2 × 106/ml) were incubated in a shaking water bath at 37°C before addition of the (priming) agents CytB (5 μg/ml, 5 min) or PAF (1 μmol/L, 2 min) and further stimulation with fMLP (1 μmol/L). After the incubation time, the CXCR3 surface expression was analyzed by flow cytometry. Green line histogram: isotype control; blue-filled histogram: CXCR3 surface expression. C, Coexpression of CXCR3 and CD63. Neutrophils were isolated and stimulated with CytB plus fMLP or PAF plus fMLP as described above and were costained with CXCR3, CD63, and CD66b. Where indicated, isotype controls were used instead of the specific Abs.

FIGURE 9.

CXCR3 expression in neutrophils. A, Confocal microscopy. Neutrophils were isolated from peripheral blood of healthy controls (2 × 106/ml). Where indicated, neutrophils were permeabilized with an equal volume of cold 0.1% (w/v) Triton X-100. Neutrophils were stained with rabbit anti-CXCR3 Abs, anti-rabbit-Ig Alexa Fluor 555, and Con A conjugated to Alexa Fluor 488. Confocal laser-scanning microscopy was performed with the Leica TCS NT laser system using a Leica DM IRB microscope. Red color represents CXCR3 receptors and green color Con A. B, Mobilization of intracellular CXCR3 expression. Neutrophils (2 × 106/ml) were incubated in a shaking water bath at 37°C before addition of the (priming) agents CytB (5 μg/ml, 5 min) or PAF (1 μmol/L, 2 min) and further stimulation with fMLP (1 μmol/L). After the incubation time, the CXCR3 surface expression was analyzed by flow cytometry. Green line histogram: isotype control; blue-filled histogram: CXCR3 surface expression. C, Coexpression of CXCR3 and CD63. Neutrophils were isolated and stimulated with CytB plus fMLP or PAF plus fMLP as described above and were costained with CXCR3, CD63, and CD66b. Where indicated, isotype controls were used instead of the specific Abs.

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The present study provides evidence in humans that the inflammatory microenvironment induces a novel chemokine receptor repertoire on infiltrated neutrophils that expands their functional responsiveness to surrounding chemokines. Distinct mechanisms were found to induce chemokine receptor expression on neutrophils, mediated via TLR/NOD2 activation, cytokine stimulation, or neutrophil senescence/apoptosis. After induction of chemokine receptors in chronic lung diseases in situ, activation of the receptors differentially modulated neutrophil effector responses. Since the up-regulation of chemokine receptors was exclusively found on infiltrated neutrophils at inflammatory sites, the targeting of these chemokine receptors may have the potential to site-specifically target neutrophilic inflammation.

In contrast to eosinophils, human neutrophils are traditionally known to express a very restricted pattern of CXC chemokine receptors (13) and are unresponsive to CC chemokines (6, 7). Data obtained from animal models, however, suggest that under inflammatory conditions neutrophils are able to respond to chemokines that are functionally inactive under resting conditions and express chemokine receptors that are absent on quiescent neutrophils (14, 18, 43, 44, 45, 46). In particular, CCL2 was found to elicit neutrophil transendothelial migration in adjuvant-induced vasculitis in rats, whereas neutrophils from naive animals did not respond to the chemokine (18). Consistent with the functional response, increased expression levels of CCR1 and CCR2 were detected on neutrophils from adjuvant-immunized animals, while both receptors were absent on naive neutrophils. The potential role of CC chemokine receptors in neutrophil recruitment is further corroborated by the finding that murine neutrophils during sepsis, following cecal ligation and puncture, bind and respond to CCL2 and CCL3, associated with increased mRNA expressions of CCR1, CCR2, and CCR5 in neutrophils (44). In a murine ischemia-reperfusion model, genetic knockout of CCR1, CCR2, or CCR5 impaired neutrophil migration to postischemic tissue through effects on intravascular adherence and transmigration (47). Neutralization of CCL3 in murine models of LPS-induced lung injury attenuated pulmonary neutrophil influx (45, 46), and neutrophils from inflammatory exudates migrated to CCL3 (43). Previous studies with peripheral blood-isolated human neutrophils found that GM-CSF and TNF-α primed neutrophils for migration to CCL3 via a CCR5-dependent mechanism (19, 21). However, the expression and functional role of chemokine receptors on infiltrated neutrophils in human inflammatory diseases has not been investigated so far. Our data provide evidence that neutrophils at the pulmonary and synovial site of inflammation express CCR1, CCR2, CCR3, CCR5, CXCR3, and CXCR4, which are absent or only marginally expressed on peripheral blood neutrophils. The expression pattern of these chemokine receptors was inversely related to the expression characteristics found for conventional CXCR1 and CXCR2 receptors on neutrophils, which were highly expressed on circulating cells but were decreased on infiltrated neutrophils. Although CXCR2+ neutrophils were less frequently found in BALF/SF compared with peripheral blood in general, CXCR1+ neutrophils were more specifically reduced in BALF from patients with chronic lung diseases compared with BALF from healthy controls, which is in line with previously published results on CXCR1/CXCR2 (48).

Since the expression of these chemokine receptors in situ was associated with high amounts of the respective chemokine ligands in the inflammatory microenvironment, a physical interaction of these chemokines with chemokine receptor-expressing neutrophils is probable. This interaction has functional consequences, since we found that infiltrated neutrophils are capable of responding to these chemokines. Chemokine receptor-expressing neutrophils isolated from BALF or SF of patients with chronic lung diseases or RA migrated to their respective chemokine ligands ex vivo. Because the migratory capacity of these isolated cells was relatively low, we performed studies to discriminate chemotaxis from chemokinesis. These studies confirmed that the observed chemokine-induced cell migration was indeed gradient-dependent chemotaxis. The ligand activation of chemokine receptor-expressing neutrophils isolated from BALF of patients with COPD resulted in modulation of antibacterial neutrophil effector responses. Specifically, the proinflammatory chemokines CCL2, CCL3, and CXCL11 enhanced antibacterial neutrophil responses, whereas the homeostatic chemokine CXCL12 suppressed fMLP-induced respiratory burst and bacterial killing capacity by neutrophils. Regarding the complex situation in vivo, chemokines produced at sites of inflammation may have a synergistic effect on local neutrophil responses as shown previously for the cooperation between IL-8 and other chemokines in vitro (49).

To unravel which factors induce the novel chemokine receptor expression on infiltrated neutrophils, we mimicked in vitro several conditions characteristic for the inflammatory microenvironment, in particular proinflammatory cytokines, long-term survival, and contact with pathogens resulting in activation of TLR/NOD receptors on neutrophils. These studies demonstrated that CCR1, CCR3, CCR5, CXCR3, and CXCR4 surface expressions were induced via different mechanisms. CCR1 and CCR3 were up-regulated through cytokines, CCR5 via NOD2 activation and apoptosis, CXCR3 through TLR1/TLR2 activation, and CXCR4 in part via apoptosis. GM-CSF and IFN-γ up-regulated CCR1- and IFN-γ-induced CCR3 expression, which is in line with two previous studies showing that GM-CSF and IFN-γ induce mRNA expression of CCR1 and CCR3 in neutrophils in vitro (8, 20). We extend these findings by showing an effect of these cytokines on protein levels of CCR1 and CCR3 and further demonstrate that the combination of GM-CSF, TNF-α, and IFN-γ is the most potent inducer of CCR1 and CCR3 expression. In contrast to all other chemokine receptors analyzed, CXCR4 was down-regulated by proinflammatory cytokine treatment. This finding is consistent with previous reports by Bruhl et al. (50) and Nagase et al. (51) demonstrating that CXCR4 on neutrophils is internalized through the action of proinflammatory cytokines. We found that long-term culture up-regulated CCR5 and CXCR4 surface expression in a protein synthesis-dependent manner, whereas no effect on CCR1, CCR2, CCR3, or CXCR3 was observed. Interestingly, after 24 h of culture, CCR5+ neutrophils were mainly annexin V+, while CXCR4+ neutrophils were only partially apoptotic. In striking contrast, CCR5+ and CXCR4+ infiltrated neutrophils in BALF were almost completely annexin V, suggesting that factors in vivo preserve neutrophil viability but allow the up-regulation of CCR5 and CXCR4. G-CSF and GM-CSF are well known to promote neutrophil survival (52, 53). Given this fact, we speculate that the high levels of G-CSF and GM-CSF present in BALF of patients with chronic lung diseases (54) may protect neutrophils from apoptosis in situ.

CCR5 has been found previously to be expressed on apoptotic murine neutrophils and to play a role in sequestering CCR5 ligands during resolution of inflammation in murine peritonitis (55). This finding in mice is in line with our observation in human neutrophils that CCR5 expression was induced by apoptosis in vitro. The functional significance of CCR5 on neutrophils remains to be established, but based on our findings and the data from the latter murine study, we speculate that CCR5, induced on human neutrophils upon apoptosis, may similarly sequester CCR5 ligands in situ. Several pieces of evidence in our data, however, suggest that apoptosis may not be the only factor that induces and regulates CCR5 and CXCR4 expression on infiltrated human neutrophils: 1) CCR5 and CXCR4 expressions were mainly found on nonapoptotic BALF neutrophils in situ; 2) infiltrated pulmonary neutrophils, in contrast to in vitro generated apoptotic neutrophils (55, 56), were capable of migrating to CCR5 (CCL4) and CXCR4 (CXCL12) ligands, which is in line with previous reports on neutrophil migration to CCR5 (21) and CXCR4 (50) ligands; and 3) TLR1/2/NOD2 activation induced CCR5 surface expression independently from apoptosis. To further elucidate the effect of the pulmonary microenvironment on chemokine receptor expression on neutrophils, we treated isolated neutrophils with several TLR ligands that are abundantly present in chronic bacterial lung diseases such as CF and COPD lung disease (41). These studies showed that CCR5 and CXCR3 up-regulation were susceptible toward TLR/NOD2 receptor activation. Although TLR2/NOD2 ligands (57) induced TLR5 surface expression, TLR1/TLR2 activation strongly up-regulated CXCR3 surface expression on neutrophils. These findings suggest that the encounter of neutrophils with bacterial pathogens modulates their chemokine receptor expression pattern.

CXCR3, which shares an amino acid homology with CXCR1 and CXCR2 of 50–60% (58), is traditionally known to be expressed on activated Th1 lymphocytes associated with chronic inflammatory diseases and has not been described to be expressed on neutrophils so far (59). Our study demonstrates that CXCR3 surface expression was present on infiltrated neutrophils in inflammatory human diseases in situ. We further found that CXCR3 was stored in intracellular granules in resting neutrophils and exocytosis experiments demonstrated that the appearance of CXCR3 surface expression was paralleled by translocation of intracellular CD63+ neutrophil granules to the cell surface. Together with the fact that CytB plus fMLP, but not PAF plus fMLP, up-regulated CXCR3 surface expression, these findings indicate that CXCR3 is stored in azurophil granules in neutrophils. This is the first demonstration (to the best of our knowledge) that neutrophils store a chemokine receptor in primary granules.

In contrast to inflammatory chemokine receptors, the homeostatic chemokine receptor CXCR4 regulates basal trafficking of immune cells (60). Inducible surface expression of CXCR4 on human neutrophils has been described previously (50, 51, 61), and neutrophils in SF from patients with inflammatory joint diseases were CXCR4+ (62), which is consistent with our findings. High levels of the CXCR4 ligand CXCL12 have been found in SF from RA patients (63), where CXCL12 is suggested to contribute to the accumulation of leukocytes (64, 65). In vitro, CXCL12 was found to mediate desensitization of SF-/fMLP-activated respiratory burst in neutrophils (66). The role of the CXCL12-CXCR4 axis in lung diseases is less clear. Our study is the first to show that infiltrated neutrophils in chronic lung diseases highly express CXCR4 on their surface and are surrounded by high levels of CXCL12 in the airway microenvironment. We further show that CXCL12 via CXCR4 dampens the respiratory burst activity by pulmonary neutrophils. Further evidence on the role of CXCR4 in lung disease comes from a study in a mouse model of allergic asthma, demonstrating a beneficial effect of the CXCR4 antagonist AMD3100 on pulmonary inflammation (67). Another murine study found that senescent CXCR4high neutrophils preferentially home to the bone marrow in a CXCR4-dependent manner, suggesting a role for CXCR4 on neutrophils for the clearance of senescent neutrophils (61). Despite these intriguing findings, further studies are required to fully understand the functional significance of CXCR4 on infiltrated neutrophils in chronic inflammatory diseases.

In summary, our data indicate that infiltrating neutrophils acquire a novel chemokine receptor expression repertoire and functional chemokine responsiveness, enabling them to adapt to chronic inflammatory conditions. These data demonstrate that chemokine receptors on neutrophils are not only involved in cell migration, but specifically modulate cellular effector functions at the site of inflammation. Inducible chemokine receptors on infiltrated neutrophils provide a novel site-specific target to modulate neutrophilic inflammation in chronic lung disease.

We thank Andrew D. Luster and Benjamin D. Medoff, Center for Immunology and Inflammatory Diseases, Massachusetts General Hospital, Harvard University, for helpful discussions. We thank Nikolaus Rieber, University Childrens’ Hospital, Munich, Germany, for critical revisions of this manuscript.

The authors have no financial conflict of interest.

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.

2

Abbreviations used in this paper: BALF, bronchoalveolar lavage fluid; SF, synovial fluid; RA, rheumatoid arthritis; PAF, platelet-activating factor; CHX. cycloheximide; LTA, lipoteichoic acid; poly(I:C), polyinosine-polycytidylic acid; PGN, peptidoglycan; MDP, muranyl peptide; Pam3CSK4, Pam3CysSerLys4; CF, cystic fibrosis; COPD, cystic obstructive pulmonary disease; FEV1, forced expiratory volume in 1 s; MFI, mean fluorescence intensity; PI, propidium iodide; FSC, forward scatter; SSC, side scatter; LY, Lucifer Yellow; DHR, dihydrorhodamine 123; CytB, cytochalasin B.

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