Eosinophils and monocytes produce pulmonary and activation-regulated chemokine, which activates cultured monocytes/macrophages

Ingrid Schraufstatter, Hiroshi Takamori, Lyudmila Sikora, P. Sriramarao, Richard G. DiScipio


Pulmonary and activation-regulated chemokine (PARC/CCL18) belongs to the family of CC chemokines and shares 61% sequence identity with monocyte inflammatory protein (MIP)-1α. Produced by dendritic cells and macrophages primarily in the lung, PARC is known to be chemotactic for T cells. Because PARC's biological function is largely unknown, we screened various leukocyte populations for PARC expression and for response to PARC, with the idea that the cellular source may link PARC to disease states in which it may be involved. Here we report that eosinophils obtained from individuals with mild eosinophilia express PARC as assessed by RT-PCR on eosinophil RNA. The eosinophil preparations were free of monocytes, a known source of PARC, and no RT-PCR product was obtained from neutrophils. Furthermore, PARC protein was detected by ELISA in the supernatants of eosinophils from seven of nine donors and in higher concentration in the supernatants of monocytes on day 1 of culture. Purified recombinant PARC activated human monocytes/macrophages kept in culture for 3-4 days but not freshly isolated monocytes. The threshold dose for Ca2+ mobilization as determined fluorometrically in indo 1-AM-labeled monocytes was 5 nM; maximal response was reached with ∼50 nM PARC. PARC was chemotactic for these cultured monocytes and caused actin polymerization determined by FITC-phalloidin binding and fluorescence-activated cell sorting analysis. In contrast, PARC activated neither neutrophils nor eosinophils. Eosinophil production of PARC, its chemotactic effect on monocytes and lymphocytes, and PARC's previously described localization to the lung suggest that this chemokine might play a role in pulmonary leukocyte trafficking.

  • novel source
  • novel target cell

pulmonary and activation-regulated chemokine (PARC/CCL18), also known as DC-CK-1, AMAC-1, monocyte inflammatory protein (MIP-4), and SCY A 18, is a member of the CC chemokine family, and most closely related to MIP-1α with which it shares 61% sequence identity (1, 16). Despite this high homology to MIP-1α, PARC does not activate CC chemokine receptor (CCR) 1 to CCR8 (26), and so far no receptor has been defined that is activated by PARC (16, 21, 23). However, Nibbs et al. (21) recently reported that PARC antagonizes CCR3 and competes with eotaxin, implying that PARC may be an inhibitory cytokine for certain cell types. Because the CCR3 is most prominently expressed by eosinophils, the question arose whether these cells may regulate a response to eotaxin through PARC expression.

Expression of the mRNA for PARC has been detected in human lung, particularly in alveolar macrophages, in follicular dendritic cells in the germinal centers of lymph nodes, in peripheral blood monocytes following stimulation with lipopolysaccharide (LPS), and in low concentrations in bone marrow (16). It has also been detected by in situ hybridization in atherosclerotic plaque (27). The known sources of PARC, primarily lung macrophages, give little information about its possible functions. Because PARC has been detected only in primates to this point, insights into its function from animal studies have been limited (13). However, in a recent gene array study in a monkey model of allergic asthma, PARC, along with eotaxin and monocyte chemotactic protein (MCP)-1, was one of the most highly induced genes in the lung tissue following allergen challenge (35), suggesting a role for PARC in allergic inflammation. Therefore, we systematically investigated which leukocytes, in particular those associated with allergic inflammation, were capable of producing PARC. Apart from monocytes, we found that eosinophils produce PARC, which was detected both at the mRNA level and at the protein level in the supernatants of peripheral blood eosinophils.

The cellular responses caused by PARC have not been investigated thoroughly. It has been shown that PARC is a chemotactic factor for various subclasses of lymphocytes in vitro (1) and in vivo (13). When PARC was injected into mice, only a moderate number of lymphocytes accumulated at the site of administration (13). However, since it is doubtful that mice express a PARC equivalent (31), and chemokine/chemokine receptor usage in rodents is quite different from that seen in humans, the lymphocyte-specific response observed in mice might be only a partial presentation of the human response. For this reason, we searched for additional cells that could respond to PARC and found that peripheral blood monocyte/macrophages in culture for 3-4 days respond to recombinantly expressed PARC. Freshly isolated monocytes did not show this activity (16). This distinction between freshly isolated monocytes and monocyte/macrophages may help in the identification of the PARC receptor.


Reagents. The clone for PARC from a human lung cDNA library (Stratagene, La Jolla, CA) was a gift of Dr. J. Han (The Scripps Research Institute). The DNA was amplified by 30 cycles of PCR using the forward 5′-oligonucleotide GAATTAATGCACAAGTTGGTACCAAC and the reverse 3′-oligonucleotide GGAGGATCCAGGCATTCAGCTTCAG and PCR reagents from GIBCO-BRL Life Sciences (Grand Island, NY). The PCR product was cloned into the pBAD TOPO TA cloning vector (Invitrogen, Carlsbad, CA), excised with VspI and BamHI, and subcloned into the NdeI/BamHI cut bacterial expression vector pET-15b (Novagen, San Diego, CA), which adds an amino-terminal thrombin-cleavable oligohistidine tag to the expressed protein. The correctness of the gene product was verified by automated DNA sequencing (Scripps Core Facility, La Jolla, CA).

For cDNA preparation RNA was purified with the RNeasy kit (Qiagen, Valencia, CA) followed by reverse transcription with the Promega RT-PCR kit (Promega, Madison, WI) according to the manufacturer's recommendations. Possible genomic DNA contamination could be excluded since the PARC PCR primers would have included an intron region in the genome.

MIP-1α, an ELISA kit for MIP-1α, and neutralizing antibodies against the IL-8 receptor 1 (CXCR1) and CXCR2 were purchased from R&D Systems (Minneapolis, MN). IL-8 and growth-related protein (GRO)-α were expressed recombinantly and purified as previously described (29). LPS (Re595 Salmonella minnesota) was from Sigma Chemical (St. Louis, MO), and tumor necrosis factor-α (TNF-α) from Biosource (Sunnyvale, CA). Indo 1-AM and FITC-phalloidin were obtained from Molecular Probes (Eugene, OR).

Purification of PARC. Recombinant PARC expression in Escherichia coli strain HMS 174(DE3) (Novagen) was induced with 0.1 mM isopropylthiogalactoside for 3 h at 37°C. After centrifugation, the bacterial pellet was suspended in 20 mM sodium acetate, pH 5.0/0.15 M NaCl/5 mM EDTA/10 mM benzamidine/20 mM ϵ-amino caproic acid/0.2% Triton X-100, sonicated for 30 s, frozen, and thawed. After centrifugation, the pellet was dissolved in 8 M urea, 10 mM HEPES, pH 6.5, and 0.15 M NaCl, and the sample was absorbed to and eluted from Tris(carboxymethyl)ethylinediamine-Sepharose, which had been charged with NiCl2 (24). Expressed PARC was eluted with 40 mM sodium acetate, pH 4.5, 6 M guanidine HCl, and dialyzed against 10 mM HEPES/0.15 M NaCl, pH 7.0. The oligohistidine leash was excised from the expressed polypeptide by incubation with thrombin for 15 h at 23°C using an enzyme to substrate weight ratio of 1:4,000. After inactivation of the thrombin with 0.1 mM PMSF, PARC was separated from contaminating polypeptides by cation exchange chromatography on a dextran sulfate Sepharose column with a linear salt gradient from 0.15-1.5 M NaCl. Matrix-assisted laser desorption-time of flight mass spectroscopy of isolated PARC was performed as a service at The Scripps Research Institute.

Immunological detection of PARC. Polyclonal antibodies to recombinant PARC were raised in rabbits by Strategic Biosolutions (Ramona, CA), and a sandwich ELISA was developed with this antibody preparation. The rabbit IgG was affinity purified on recombinant PARC covalently bound to Sepharose using the cyanogen bromide reaction, and the purified antibody was biotinylated using NHS-LC-biotin (succinimidyl-6-biotinamide-hexanoate; Pierce Chemical, Rockford, IL). Polyclonal antibody (10 μg/ml) was absorbed to microtiter wells (Nunc maxisorp; Nalge Nunc International, Rochester, NY) for 16 h in 150 mM sodium carbonate, pH 9.6. In later experiments monoclonal anti-PARC (2 μg/ml, R&D Research) was used, resulting in slightly increased signal-to-noise ratio. Each plate was washed 3× with 20 mM imidazole HCl, pH 7.3/0.15 M NaCl, blocked with 2% BSA/0.5% Tween 20 in 0.1 M sodium phosphate, pH 7.0/0.15 M NaCl for 1 h and washed. Cell supernatants or PARC standards in Ultraculture (Bio-Whittaker, Walkersville, MD) containing 1 mg/ml BSA and 1 mg/ml ovalbumin were added and incubated for 2 h at room temperature. After three further washes, 100 μl of biotinyl-anti-PARC-IgG were added, and the plate was incubated for 1 h at room temperature. The plate was again washed, incubated with avidin-horseradish peroxidase conjugate (DAKO, Carpinteria, CA), and finally developed with 2-2′-azino-di-3-ethylbenzazoline sulfonate (Roche Molecular Biochemicals, Indianapolis, IN), at 0.5 mg/ml in 50 mM sodium phosphate/20 mM sodium citrate, pH 5/0.03% H2O2, and the absorbance was read at 405 nm on a Molecular Devices Vmax reader (Molecular Devices, Sunnyvale, CA). All samples were analyzed at least in duplicate from cell preparations from 11 donors. The ELISA was specific for PARC and did not cross-react with human MIP-1α or MIP-1β, the two most closely related chemokines. Nor was there cross-reactivity with regulated on activation normal T-cell expressed and presumably secreted (RANTES), MCP-1, eotaxin, IL-8, or GRO-α.

Preparation of leukocytes. Mononuclear and neutrophil fractions were purified from fresh human acid citrate dextrose blood from healthy donors by Hetastarch sedimentation followed by Percoll gradient centrifugation (34) (Amersham Pharmacia Biotech, Piscataway, NJ). The neutrophil fraction was >98% pure. We further separated the mononuclear fraction into monocytes and lymphocytes by allowing the monocytes to adhere to tissue culture plastic in RPMI 1640 containing 10% FCS for 90 min at 37°C. Nonadherent lymphocytes in the supernatant were then removed, and the remaining monocytes were thoroughly washed and cultured for 1-4 days in RPMI 1640 containing 10% FCS (GIBCO-BRL). For calcium mobilization and actin polymerization assays, adherent monocytes were removed from the surface by a 10-min incubation with cell dissociation buffer (GIBCO-BRL), and the nonadherent fraction was added. The combined cells were washed and resuspended in Hanks' balanced salt solution (HBSS; 140 mM NaCl, 5 mM KCl, 3.8 mM KH2PO4, 2.2 mM Na2HPO4, and 5.5 mM glucose, pH 7.4). For chemotaxis, only the nonadherent cells were used to avoid possible inactivation of adhesion molecules due to cell processing. Overnight cultures of monocytes used in the ELISA were maintained in Ultraculture (Bio-Whittaker). Fluorescence-activated cell sorting (FACS) analysis showed that these cells were between 94 and 98% monocytes and 2 and 6% lymphocytes as assessed by size and granularity and by determining the percentage of CD14-positive cells using anti-CD14 (M5E2 monoclonal; Pharmingen, La Jolla, CA) followed by staining with FITC-anti-mouse IgG (Biosource, Camarillo, CA).

Eosinophils were isolated from blood of healthy, mildly allergic donors. After the preparation of the polymorphonuclear cell fraction as described above, eosinophils were purified by negative selection using anti-CD16 with magnetic bead separation (14). The eosinophil preparations were >99% pure with neutrophils as the only trace contaminating cells. Mononuclear cells were never observed in eosinophil preparations as assessed by hematoxylin/eosin Y staining. When determining the release of PARC from eosinophils, cells were incubated overnight in Ultraculture with the addition of 5 ng/ml of IL-5 and granulocyte-macrophage colony-stimulating factor (GM-CSF) (both from R&D Systems).

All leukocytic cell lines were obtained from ATCC and grown in RPMI 1640 containing 10% FCS.

Calcium mobilization. To detect changes in intracellular calcium, we labeled leukocytes for 30 min with indo 1-AM (Molecular Probes) as previously described (29). Fluorescently labeled cells (2 × 105) were warmed to 37°C for 2 min in a stirred cuvette containing 200 μl of modified Gey's buffer (MGB; 140 mM NaCl, 5 mM KCl, 1.9 mM KH2PO4, 0.5 mM MgCl2, 1.1 mM Na2HPO4, 1.5 mM Ca2Cl, 10 mM HEPES, and 5.5 mM glucose, pH 7.4), the stimulus was added, and the emission ratio at 400/480 nm was followed kinetically on an SLM 8000 fluorometer (SLM Aminco, Rochester, NY) as described previously (32). Calcium concentrations were calculated as described in Ref. 12.

Actin polymerization. For the determination of actin polymerization, 4-day-old monocyte/macrophages or lymphocytes were stimulated with PARC for the indicated time at 37°C and pipetted into a mixture of formaldehyde, lysophosphatidic acid, and FITC-phalloidin (Sigma Chemical) as previously described (7). The mean fluorescence of the cell population was detected by FACSCAN analysis (Becton Dickinson, Research Triangle Park, NC) of the monocyte or lymphocyte population that was selected by its characteristic size and granularity.

Chemotaxis assay. Monocytes (2 × 105) in HBSS containing 0.5% BSA were applied to the top compartment of 5-μm pore-size Transwell filters (Costar; Corning, Acton, MA) and allowed to transmigrate into the bottom compartment containing varying concentrations of PARC for 90 min at 37°C. At this time, 5 mM EDTA was added to the lower compartment to release monocytes that adhered to the underside of the filters, and transmigrated cells were counted manually in 25 low-power magnification fields. In some experiments the filters were coated with 200 μl/ml Matrigel (Becton Dickinson, San Diego, CA) overnight at 4°C and washed three times with 150 mM NaCl and 10 mM HEPES, pH 7.3, to mimic basement membrane-like conditions before the addition of monocytes.


Expression of PARC in E. coli. PARC was originally cloned from a human lung cDNA library. The coding sequence for the mature protein was amplified by PCR and cloned into the pET-15b bacterial expression vector. Figure 1 depicts an SDS gel of the purified recombinant protein, in which lane 1 shows the propeptide still containing the histidine leash and lane 2 the thrombin-cleaved protein used in all of the experiments. Due to its high content of basic amino acids (10 out of 69), PARC shows a slightly slower mobility on SDS-PAGE than the calculated molecular weight. Matrix-assisted laser desorption-time of flight mass spectrometry indicated a mass for unmodified recombinant PARC of 8,246, exactly matching the calculated mass of PARC after factoring in four additional amino acids at the NH2 terminus and the loss of four hydrogen atoms after formation of two disulfide bridges.

Fig. 1.

Purification of pulmonary and activation-regulated chemokine (PARC) from Escherichia coli. PARC was purified from E. coli as described in materials and methods and applied to a gradient SDS-polyacrylamide gel. Lane 1: oro-PARC retaining the histidine leash; lane 2: PARC after thrombin cleavage of the histidine leash; lane 3: molecular weight (MW) standards: carbonic anhydrase (MW 29,000), soybean trypsin inhibitor (MW 21,000), pancreatic RNase (MW 14,000), and C3a (MW 8,800).

Expression of PARC mRNA by various leukocytes. Screening for expression of PARC in various tissues indicates that PARC mRNA was not limited to the lung but was produced by various cells of leukopoietic lineage. RT-PCR indicates that it was expressed in bone marrow, by peripheral monocytes following 90 min of adhesion to plastic, and by freshly isolated eosinophils (Fig. 2), but not by neutrophils. The trace of a PCR product observed in lymphocytes may have been caused by monocyte contamination. A PCR product for PARC was obtained from the eosinophil mRNA of three consecutive donors (Fig. 2), and DNA sequencing confirmed that the eosinophil-derived PARC PCR product was identical to the recombinant PARC derived from lung cDNA.

Fig. 2.

RT-PCR of PARC from human lung, monocytes, nonadherent leukocytes, neutrophils, bone marrow, and eosinophils from 3 different donors. A: lane 1, lung cDNA (Stratagene); lane 2, bone marrow cDNA (Stratagene); lane 3, monocyte cDNA prepared from adherent, freshly isolated monocytes; lane 4, peripheral lymphocyte cDNA; lane 5, neutrophil cDNA; lane 6, 1-kb DNA ladder. B: lane 1, PCR of PARC in pET 15b as a positive control; lanes 2-4, eosinophil cDNA from 3 different donors; lane 5, DNA ladder (GIBCO-BRL).

Detection of PARC by ELISA. A sandwich ELISA was established to determine PARC protein concentrations in cellular supernatants. Consistent with the PCR results, both eosinophils and monocytes expressed PARC protein constitutively. Three out of the 11 eosinophil donors presented with peripheral eosinophil counts in the normal range (<3% of the leukocyte population). In contrast to the eight eosinophilic donors, PARC levels in these three donors were below the detection limit of the ELISA. The large variation in PARC concentrations observed with different donors (290+/-200 pg/106 eosinophils and 760+/-420 pg/106 monocytes; Fig. 3, A and B) is not unusual for the expression of a chemokine. Although eosinophils produced only a third of the amount of PARC obtained from monocytes, this may represent a sizable contribution under conditions of allergic inflammation, where tissue eosinophils can be the dominating cell population.

Fig. 3.

PARC protein expression by eosinophils (eos.) and monocytes (MΦ). PARC and MIP-1α concentrations were determined by sandwich ELISA as described in materials and methods. A and B: concentration of PARC in the supernatants of eosinophils, neutrophils (PMNs, A), and monocytes (B) (1.5 × 106 cells/ml), and of monocyte inflammatory protein (MIP)-1α in monocytes (B). Eosinophils and monocytes were cultured for 16 h; PMNs, because of their limited viability, for 2 h. Eosinophil viability was optimized by incubating the cells with IL-5 and granulocyte-macrophage colony-stimulating factor (5 ng/ml each). Each data point represents the mean of triplicate ELISA determinations of the leukocyte preparations from separate donors. The horizontal line indicates the threshold of sensitivity for the ELISA. C: effect of various stimuli on PARC production by monocytes incubated for 16 h at 37°C. The stimuli were 100 ng/ml LPS, 10-8 M IL-8, and 10-8 M growth-related protein (GRO)-α, as indicated underneath each bar (means ± SD, n = 6-13). Neutralizing antibody against the IL-8 receptor 1 (CXCR1) or CXCR2 was added at 10 μg/ml where indicated. D: effect of various stimuli on MIP-1α production by monocytes incubated for 16 h at 37°C. The stimuli were 100 ng/ml LPS or 10-8 M IL-8, as indicated underneath each bar (means ± SD, n = 5). Neutralizing antibody against the CXCR1 or CXCR2 was added at 10 μg/ml where indicated.

PARC concentrations were comparable to MIP-1α concentrations of the same monocyte supernatants, both in terms of concentration and variability (620+/-390 pg/106 cells, Fig. 3B). It should also be noted that our method of monocyte purification by means of their adhesion to plastic has been found to activate the release of a number of cytokines and chemokines without the need for inflammatory mediators (17, 30). In this sense these cells are not strictly unstimulated but stimulated in a fashion that mimics monocyte extravasation in vivo.

No PARC was detected in the supernatants of neutrophils or lymphocytes or in monocyte/macrophages kept in culture for 4 days. LPS increased protein expression of PARC by monocytes (Fig. 3C) as previously reported for PARC mRNA levels (16). Again, the increase of PARC in the supernatants of LPS-stimulated cells was in the same concentration range as that observed for MIP-1α (Fig. 3D). IL-8 also increased expression of PARC to a statistically significant degree (Fig. 3C; Student's t-test, P < 0.05). In contrast, GRO-α failed to induce increased PARC expression, suggesting that the response to IL-8 was mediated by the CXCR1, which is specific for IL-8 (Fig. 3), and not by the CXCR2, which has similar affinities for IL-8 and GRO-α (18, 29). To test this, monocytes were incubated with anti-CXCR1 or anti-CXCR2 antibody (10 μg/ml, R&D Systems) in the presence or absence of IL-8 (10 μg/ml, R&D Systems). Anti-CXCR1, but not anti-CXCR2, antibody significantly decreased the concentration of PARC in the supernatant of IL-8-stimulated cells (P < 0.01, Fig. 3C). Interestingly, anti-CXCR1 antibody by itself reduced PARC concentrations in unstimulated cells to levels below those seen in the absence of antibody (Fig. 3C), suggesting that IL-8 produced by the monocytes, which are known to produce IL-8 under our culturing conditions, contributed to the basal PARC production in these cells.

Screening for cell lines responding to PARC. Because calcium mobilization is a facile assay that allows fast screening of many different cell types and uses small cell numbers (2 × 105 cells/sample), we used this assay to assess cell types that express receptors for PARC. Numerous leukocytic cell lines failed to respond to PARC. These included THP1, HL60, U937, Ramos, Raji, Jurkat, and RBL2H3 cells and included differentiation with PMA for HL60 and THP1 cells, with dibutyryl-cAMP for HL60 cells and U937 cells and interferon-γ and G-CSF for U937 cells. Among primary leukocytes, only lymphocytes and 3- to 4-day-old monocytes (Fig. 4) responded to PARC with calcium mobilization. Eosinophils and neutrophils showed no response.

Fig. 4.

Calcium mobilization in monocytes stimulated with PARC. Dose response and comparison between different activators. Four-day-old monocyte/macrophages were labeled with indo 1-AM, and 2 × 105 cells in 200 μl of modified Gey's medium were stirred at 37°C, and the fluorescence ratio at 400/480 nm was determined kinetically following the addition of stimulus. A: tracings of the fluorescence ratio over time with 3 concentrations of PARC, which was injected at 10 s (arrow). At the end of 1 determination 0.1% Triton X-100 was added to determine the total indo 1-labeled intracellular Ca2+ (shown in 1 of the samples). B: monocyte/macrophages on day 4 were stimulated with 5 different stimuli (PARC, open bar; MIP-1α, stippled bar; IL-8, filled bar; GRO-α, horizontal stripes; C3a, gray bar). At this time the monocytes responded to all 4 chemokines with slightly higher sensitivity for MIP-1α, IL-8, and GRO-α than for PARC. No response was seen to C3a. The maximal calcium response was defined as the peak of the Triton X-100-induced calcium mobilization. Means of duplicate determinations for each condition. One experiment representative of 4.

Calcium mobilization in monocytes/macrophages. Because lymphocyte stimulation by PARC had been described previously (1, 16), we primarily investigated the effect of PARC on monocytes. In 4-day-old monocytes/macrophages, PARC induced a calcium flux with a threshold dose of ∼5 × 10-9 M, and the maximal response, corresponding to a rise of intracellular Ca2+ from 140 nM to 1 μM, was reached with 10-7 M PARC (Fig. 4). To assure that the Ca2+ flux could not be attributed to contaminating lymphocytes, 2 × 104 lymphocytes, a cell number that is greater than the lymphocyte contamination in any of our monocyte preparations, were stimulated with PARC. Under these conditions no change in calcium fluorescence was observed, indicating that monocytes were the cells that responded to PARC.

Monocytes purified as the adherent cell fraction in the mononuclear cell mixture failed to respond to PARC, when used on the day of cell preparation. This refractory behavior of freshly isolated, adherent monocytes was not surprising, since these cells expressed mRNA for PARC and secreted PARC protein as shown above, which is expected to desensitize and/or downregulate any PARC receptors these cells may possess. The same behavior was observed for several other chemokines (IL-8 and GRO-α), which like PARC failed to activate monocytes on days 0-2 but caused a calcium flux on days 3-4 (Fig. 4), which decreased again over the following days and became negative by 1 wk. The potency of PARC was slightly lower than that of MIP-1α, IL-8, or GRO-α (Fig. 4).

Actin polymerization in monocytes and lymphocytes. To confirm that monocytes responded to PARC, filamentous actin (F-actin) was determined in monocyte/macrophage and lymphocyte populations after 4 days in culture. As shown in Fig. 5, PARC caused an increase in F-actin content in both cell types. These experiments clearly indicate that 4-day-old monocytes/macrophages respond to PARC. Lymphocytes similarly responded with actin polymerization. Their unusually high F-actin content (the resting cell level of fluorescence intensity seen in FITC phalloidin-stained lymphocytes was sixfold higher than that in the monocyte population) further guaranteed that all lymphocytes were excluded from the monocyte population. This further showed that the actin polymerization in monocytes could not be attributed to contaminating lymphocytes.

Fig. 5.

Actin polymerization in monocytes and lymphocytes stimulated with PARC. Actin polymerization was measured by fluorescence-activated cell sorting analysis of FITC phalloidin-stained cells as described in materials and methods. Monocyte/macrophage and lymphocyte populations were selected for their size and granularity. Preliminary testing showed that the monocyte population was 99% CD14 positive. Means + SD of samples from 4 blood preparations. Because lymphocytes show an unusually high polymerized actin content, the ordinate is plotted as the mean fluorescence channel, which clearly indicates that the monocyte response cannot be explained by contamination with lymphocytes, which have a baseline F-actin content that is 6-fold higher.

Chemotaxis of monocytes. To test whether PARC plays a role in monocyte trafficking, we determined chemotaxis of monocyte/macrophages toward PARC. As already suggested by the F-actin response, a prerequisite for chemotaxis, PARC induced chemotaxis of 4-day-old monocytes (Fig. 6) with a threshold dose of ∼2 nM and a maximal response with 20-30 nM PARC. Checkerboard analysis indicates that the cells responded with chemotaxis rather than chemokinesis. Because leukocytes have to be able to migrate through the basement membrane during the inflammatory response, PARC-dependent cell migration through Matrigel-coated filters was determined. Under these conditions PARC was a more potent chemoattractant than IL-8 in terms of both sensitivity and numbers of attracted cells (Fig. 6B). As reported previously (1), freshly isolated monocytes did not migrate toward PARC.

Fig. 6.

Chemotaxis of monocytes induced by PARC. Chemotaxis was determined as described in materials and methods. A: dose response to PARC in standard Transwell assays. Means + SD of samples from 6 different donors (n = 12). Lymphocyte contamination was <5% in any of the transmigrated samples. B: comparison between chemotaxis induced by PARC and IL-8 using Matrigel-coated Transwells. Triplicates from 1 experiment representative of 3. Underneath results are shown for checkerboard analysis to prove chemotaxis rather than chemokinesis (duplicate samples from 2 donors).

As a further functional assay in monocytes/macrophages, we determined the release of H2O2 (9) following stimulation with PARC but found no evidence for an oxidative burst (results not shown). Under the same experimental conditions, 100 ng/ml PMA caused the production of H2O2 (1.1 nmol·2 × 105 monocytes-1·60 min-1).


These results show a novel source of PARC, eosinophils, and a novel function for this chemokine, i.e., the activation of monocyte/macrophages. Previously, it had been shown that PARC mRNA is constitutively present in the lung (1, 16), primarily in alveolar macrophages, in various lymphoid tissues, and in monocytes and monocytic cell lines following stimulation with PMA (16). Here we show that eosinophils purified from peripheral blood express PARC mRNA and produce PARC protein, although PARC protein concentration could be detected in only eight out of eleven donors. The large variability between different donors is not specific for the expression of PARC but has been noted for numerous eosinophilic properties that vary considerably between different individuals (22, 25). The eight donors from whom PARC could be detected in the eosinophil supernatants all showed mild eosinophilia with eosinophil counts >400 cells/μl blood. In contrast, eosinophils amounted to <3% of the leukocyte population in the three subjects in whom the PARC concentration was below or around the detection limit of the ELISA. Generally, donors with higher eosinophil yield also showed higher PARC concentrations. Future investigation will show whether PARC expression occurs in eosinophilic subjects generally and whether there is a correlation between PARC expression and allergic conditions. PARC was detected on the mRNA level in all three subjects tested, which may reflect the increased sensitivity of PCR over the ELISA assay.

The physiological role for eosinophil-derived PARC deserves further investigation, since it may initiate both pro- and anti-inflammatory reactions. The chemotactic effect of PARC on lymphocytes (1, 13, 16) and, as shown here, on monocyte/macrophages, is likely to cause mononuclear cell attraction to an area in which eosinophils have accumulated. Thus PARC has the potential to be an important factor in pulmonary leukocyte accumulation in allergic disease. The notion that PARC may play a role in allergic lung disease is strengthened by the recent observation that PARC was one of the most strongly upregulated genes in a monkey model of allergic asthma (35).

On the other hand, PARC has an antagonistic effect on the CCR3 (21). This property of PARC may limit further eosinophil influx caused by eotaxin, RANTES, MCP-3, and MCP-4, which all stimulate CCR3 (8, 10, 23, 33). In this respect PARC may limit eosinophil accumulation by competing with the ligands that activate CCR3. Future investigation will show which of these two mechanisms, the pro- or the anti-inflammatory side, prevails and whether PARC should be inhibited or whether on the contrary PARC-derived proteins may be useful as CCR3 antagonists in asthmatics as suggested by Nibbs et al. (21).

Until now the determination of PARC expression has relied largely on its detection on the mRNA level. The development of an ELISA for PARC greatly simplifies detection of PARC and allows quantification of the expressed protein. Our results establish that monocytes express PARC also on the protein level, which increases following stimulation with LPS (Fig. 3), as suggested by the earlier detection of increased PARC mRNA levels under this condition (16, 27). In addition, IL-8 was also found to induce increased PARC expression by monocytes. In contrast, GRO-α failed to influence PARC production, suggesting that PARC induction was mediated by the CXCR1, which is specific for IL-8, and not by the CXCR2, which reacts equally well with IL-8 and GRO-α (18, 29). This was confirmed using specific antibodies. In fact baseline PARC concentrations were decreased by 50% in the presence of anti-CXCR1 antibody, suggesting that constitutive expression of IL-8 contributed to the expression of PARC by adherent monocytes. The same CXCR1-specific upregulation of monocytic chemokine production has recently been described for IL-8 itself, where IL-8, but not GRO-α, induces its own expression in an autocrine fashion (6). Eosinophils had to be cultured in the presence of IL-5 and GM-CSF to detect PARC in the supernatants. Because both these factors are, however, necessary for eosinophil survival in vitro (15), it is problematic to omit them. In addition, both IL-5 and GM-CSF are expressed by eosinophils at sites of allergic inflammation in asthmatics (5). In this respect their addition only mimics the situation seen in allergic inflammation. The concentrations of PARC found in the supernatants of eosinophils are comparable to those previously reported for IL-8 (20).

This may indicate that PARC expression was already induced in vivo in these eosinophilic subjects, but it is also possible that different stimulating agents are necessary to upregulate PARC expression in these cells.

The PARC ELISA will allow detection of this chemokine in patient samples, which is important, since the in vivo function of PARC is still unknown. Apart from its possible association with allergic disease, it may also play a role in atherogenesis, since PARC mRNA has been detected in atherosclerotic plaque (27). The detection of PARC in human fluids is essential, since its role is difficult to assess in animal models. PARC is supposed to have evolved quite recently by gene duplication from MIP-1α (13), and so far no animal equivalent of PARC has been reported. It does not appear to exist in mice (16), and its existence in rabbits is questionable. Using human PARC cDNA as a probe, we isolated five clones of rabbit MIP-1α from a rabbit lung library (Clontech) but no PARC equivalent (results not shown).

Furthermore, chemokine usage often differs between humans and rodents; for instance, mice have no IL-8 and CXCR1 equivalent but use MIP-1α for certain functions such as neutrophil chemotaxis that are mediated by IL-8 and the CXCR1 in humans. In mice, injected PARC attracted only lymphocytes and no monocytes (13). Because of the differing chemokine usage between rodents and humans, this observation does not preclude monocyte chemoattraction by PARC in humans in vivo, as would be suggested by our in vitro experiments (Fig. 6). Monocytes cultured for 3-4 days were activated by PARC as shown in calcium mobilization, actin polymerization, and chemotaxis assays. The activating concentration range was similar to that previously described for lymphocytes (16). It will be interesting to determine under which conditions monocyte/macrophages can be chemoattracted by PARC in vivo, a process that could play a role in various chronic inflammatory diseases. Because pulmonary alveolar macrophages are the most prominent source of PARC (16), PARC may contribute to the homing of monocytes to the lung.

Previous reports, which failed to show activation of monocytes by PARC (1, 16), analyzed freshly isolated monocytes, which did not respond to PARC in our hands either. This could be due either to the lack of PARC receptors at this point or to desensitization of PARC receptors from previous stimulation with cell-derived PARC. Previously, a similar refractory behavior has been observed toward IL-8 and GRO-α, which failed to stimulate monocytes that were freshly purified by means of their adhesive property, which induces IL-8 production (17). Although both classes of IL-8 receptors, the CXCR1 and the CXCR2, were shown to be present on these cells (19), no cellular response to IL-8 could be evoked in freshly prepared adhesion-isolated monocytes, and the IL-8-mediated chemotactic response of monocytes was long overlooked in vitro (11) despite evidence for it in vivo (4). A similar condition may exist for the PARC receptor on monocytes, where PARC produced by the adherent monocytes might make PARC receptors refractory to the further addition of PARC. The testing of this hypothesis will have to await the description of the PARC receptor, which is still unknown. Alternatively, PARC receptor expression may be upregulated after 3-4 days in culture.

The PARC receptor is G protein coupled (1) and sensitive to inhibition by pertussis toxin (16). This coupling to Giα is a common feature of chemokine receptors (3, 28). Although chemokine receptors can activate cellular signaling through other G proteins, coupling to Giα is necessary to induce a chemotactic response, which is mediated by the β,γ-chains of Giα (2). As already mentioned, the specific receptor activated by PARC is still unknown, but CCR1-10 have been excluded (16, 21). Our search for a cell line that responds to PARC has been similarly unsuccessful. None of the tested hematopoietic cell lines (THP1, HL60, U937, Ramos, Raji, Jurkat, and RBL2H3) was activated by PARC, which suggests that cloning of the PARC receptor has to rely on primary lymphocyte (1) or cultured monocyte/macrophage libraries, unless PARC can be shown to activate one of the existing orphan receptors.

In summary, we identified a novel source of PARC, eosinophils, and a novel function for it, the stimulation of monocyte/macrophages that were kept in culture for 3-4 days. Both these observations are of potential interest in chronic inflammatory processes.


We thank J. Han (The Scripps Research Institute) for the PARC lung cDNA.

Current address for H. Takamori: Dept. of Surgery, Kumamoto Univ., Kumamoto, Japan.


This work was supported by National Institutes of Health Grants HL-55657 (to I. Schraufstatter) and AI-35796 (to P. Sriramarao).


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