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We examined the mechanism by which lysophosphatidylcholine (LPC) regulates β2-integrin–mediated adhesion of eosiniophils. Eosinophils were isolated from blood of mildly atopic volunteers by negative immunomagnetic selection. β2-integrin–dependent adhesion of eosinophils to plated bovine serum albumin (BSA) was measured by residual eosinophil peroxidase activity. LPC caused maximal adhesion of eosinophils to plated BSA at 4 μM. Lysophosphatidylinositol, which has a similar molecular shape, mimicked the effect of LPC on eosinophil adhesion, while neither lysophosphatidylserine nor lysophosphatidylethanolamine had any effect. Phosphatidylethanolamine, a lipid that has a molecular orientation that is the inverse of LPC, blocked eosinophil adhesion caused by LPC. Unlike platelet-activating factor, a G-protein–coupled receptor agonist, LPC did not cause Ca2+-store depletion, but caused increased Ca2+ influx upon addition of Ca2+ to extracellular medium. This influx was not inhibited by U73122, a phospholipase C inhibitor, demonstrating independence from the G protein–activated phospholipase C pathway. Ca2+ influx was inhibited by either preincubation of phosphotidylethanolamine or La3+, a broad spectrum blocker of cation channels. LPC induced up-regulation of the active conformation of CD11b, which was blocked by preincubation with phosphatidylethanolamine. These data suggest that LPC causes a non–store-operated Ca2+ influx into eosinophils, which subsequently activates CD11b/CD18 to promote eosinophil adhesion.
This research focuses on the mechanisms by which lysophosphatidylcholine regulates eosinophil adhesion in vitro. This research will add to our knowledge of eosinophil infiltration in allergic inflammation, such as asthma.
Eosinophils are the predominant cells recruited to sites of inflammation in asthma and chronic allergic rhinitis (1). An initial step in this process is eosinophil adhesion to receptors on the luminal surface of vascular endothelial cells. The β2-integrin subfamily of the CD11b (Mac-1)/CD18 molcule, as well as the CD49 d (VLA-4)/CD29 of the β1-integrin subfamily, have been reported to contribute to adhesion of eosinophils to endothelial cells through binding to the endothelial ligands, intercellular adhesion molecule (ICAM)-1 and vascular cell adhesion molecule (VCAM)-1 (2). Several types of mediator that activate eosinophils and promote eosinophil adhesion to endothelium have been identified, including cytokines (such as IL-5 and granulocyte macrophage colony-stimulating factor [GM-CSF]) (3), chemokines (eotaxin, RANTES [regulated on activation, normal T cells expressed and secreted]) (4), chemoattractants (formyl-met-leu-phe [fMLP], C5a) (5), and lipid mediators (platelet-activating factor [PAF], leukotriene B4 [LTB4]) (6).
Prior reports have implicated a group of molecules with a lysophospholipid-like structure in the regulation of leukocyte trafficking (7). The mechanism of lysophospholipid-mediated cell migration has been shown to require the expression of specific G protein–coupled receptors (GPCR), such as the receptors for sphingosine 1-phosphate (S1P) (8) and lysophosphatidic acid (LPA) (9, 10). The immunologic importance of these lysophospholipid receptors is illustrated by the profound immunodeficiency observed in mice lacking expression of the S1P1 receptor in hematopoietic cells (11). S1P1-deficient T cells fail to egress from the thymus and are therefore completely absent from the periphery. The immunosuppressive drug, FTY720, has been shown to inhibit several S1P receptors, resulting in blocked egress of lymphocytes from the lymph nodes and thymus (7).
Lysophosphatidylcholine (LPC) is produced by the hydrolysis of phosphatidylcholine by various isoforms of phospholipase A2 or by reactions catalyzed by the enzyme, lecithin-cholesterol acyltransferase (12). LPC was shown to be chemotactic for monocytes and lymphocytes (13), and recently it has been reported that LPC is an apoptotic cell-derived chemotactic factor responsible for the recruitment of phagocytes to remove apoptotic debris (14). Increased expression of adhesion molecules in endothelial cells at sites of inflammation has been found to be a direct effect of LPC (15). LPC also has vasoactive properties, stimulating endothelium-dependent vascular smooth muscle relaxation (16). LPC induces oxidant generation in endothelial cells and smooth muscle cells (17), while conflicting results have been reported on human neutrophils (18, 19). While a prior study showed that LPC elicits oxidant generation in human neutrophils (18), a subsequent investigation has disputed these findings (19). LPC is found in substantial concentrations in the bloodstream (~ 200 μM ). Increased concentrations of LPC have been reported in several diseases, such as endometriosis (21), ovarian cancer (22), asthma, and rhinitis (23). In patients with asthma, the treatment with glucocorticoids resulted in an improvement in the lung function accompanied by a decrease in the plasma LPC level (24). A potential therapeutic use of LPC in sepsis (25) and cerebral ischemia (26) also has been reported. The mechanisms by which LPC protects mice against these two diseases are controversial (18, 19, 25), and prior reports (27, 28) identifying two GPCR, G2A and GPR4, as receptors for LPC now appear to be incorrect. Recent studies have found that LPC induces a change in calcium mobilization via mechanosensitive transient receptor potential channel (TRPC) 6 in cultured human corporal smooth muscle cells (29). LPC can also activate TRPC5 on vascular smooth muscle cells and HEK293 cells overexpressing this channel (30).
Inhalation of LPC to guinea pigs causes selective eosinophil infiltration in airways (31). However, the mechanism by which LPC causes eosinophil infiltration has not been defined. In this study we investigated the effect of LPC on eosinophil adhesion for β2-integrin to ICAM-1 surrogate. On the basis of preliminary data, we hypothesized that LPC causes eosinophil adhesion through activation of mechanosensitive ion channel, not GPCR. Our results demonstrate that LPC induces a non–store-operated Ca2+ influx. This effect of LPC on eosinophils, which promotes eosinophil adhesion to β2-integrin ligand bovine serum albumin (BSA), appears to be independent of GPCR normally associated with chemokines and chemoattractants.
Individual synthetic LPCs (16:0, 18:0, and 18:1 LPC), synthetic 16:0 lysophosphatidylethernolamine (LPE), synthetic 18:1 lysophosphatidylserine (LPS), lysophosphatidylinositol (LPI, 65% 16:0, 11% 18:0, and 7% 18:1) purified from soybean were obtained from Avanti Polar Lipids (Alabaster, AL). The lipids were dissolved in 50% ethanol/H2O at stock concentration of 10 mM. PAF (C16), 18:1 dioleoyl-phosphatidyl-ethanolamine (PE), and polyclonal antibody against TRPC6 were purchased from Sigma Chemical Co. (St Louis, MO). U73122 and Fura-2/AM were from Calbiochem (La Jolla, CA). Eosinophil isolation materials were obtained from StemCell Technologies (Vancouver, BC, Canada). Polystyrene 96-well microtiter plates were obtained from Costar (Cambridge, MA). Anti-CD11b mAb (clone, Bear 1) was purchased from Beckman Coulter (Fullerton, CA). The CBRM1/5 mAb against activated CD11b was a gift from Dr. T. A. Springer (Harvard Medical School, Boston, MA). Polyclonal antibody against TRPV1 was purchased from Santa Cruz (Santa Cruz, CA).
Eosinophils were isolated by a modification of the negative immunomagnetic selection technique (32). The method is based on Percoll centrifugation (density 1.089 g/ml) to isolate granulocytes, hypotonic lysis of red blood cells, and finally, immunomagnetic depletion of neutrophils by the magnetic cell separation system using anti-CD16–coated MACS particles. Eosinophil purity of 98% was routinely obtained. Cells were kept on ice until use.
β2-integrin–dependent eosinophil adhesion to plated BSA was assessed as residual eosinophil peroxidase (EPO) activity of adherent cells (5). Plated BSA has been established as a full surrogate of ICAM-1 for β2-integrin–dependent eosinophil adhesion (5, 33). Briefly, 96-well microplates were coated with 10 μg/ml BSA overnight at 4°C. The plates were washed three times with Hanks' balanced salt solution (HBSS) and saved at 4°C before use. Eosinophils (1 × 104/100 μl HBSS/0.1% gelatin) were preincubated with different concentrations of LPC for 10 min at 37°C. Cells then were added to each well of BSA-coated microplates and incubated for 30 min. After three washes with HBSS, 100 μl of HBSS/0.1% gelatin was added to the reaction wells, and serial dilutions of original cell suspension were added to the empty wells to generate a standard curve. One hundred microliters of EPO substrate (1 mM H2O2, 1 mM OPD, and 0.1% Triton X-100 in Tris buffer, pH 8.0) was then added to the wells. After 30 min incubation at room temperature, 50 μl of 4 M H2SO4 was added to stop the reaction. Absorbance was measured at 490 nM in a microplate reader (Thermomax; Molecular Devices, Menlo Park, CA). All assays were performed in duplicate. Data storage and analysis were facilitated by the use of computer software interfaced with the reader (Softmax; Molecular Devices).
Intracellular Ca2+ of eosinophils was determined by measurement of fura-2 fluorescence in a cuvette with constant stirring as described (34). In brief, cells were incubated with 2 μM Fura-2/AM for 30 min at 37°C, divided into 106 cells/tube, and kept on ice until use. Just before use, cells were warmed to 37°C and rapidly centrifuged to remove any extracellular fura-2. Cells then were transferred to a cuvette containing 2 ml HBSS without calcium and with 0.3 mM EGTA added. Fura-2 fluorescence was measured by excitation at 340 and 380 nM, with emissions recorded at 505 nM. After assessing Rmax using 10 μM ionomycin and Rmin using 5 mM EGTA, the intracellular Ca2+ was finally calculated as described by Grynkiewicz and coworkers (35).
Eosinophils were treated with various concentrations of LPC for 15 min. Thereafter, eosinophils were centrifuged at 400 × g for 10 min, and the pellets were resuspended in PBS/0.5% BSA. Aliquots of 5 × 105 eosinophils were incubated with 10 μl of fluorescein isothiocyanate–conjugated mAb against CD11b (Clone Bear 1), activated CD11b (Clone CBRM1/5), or isotype-matched control antibody for 30 min at 4°C. The cells were washed twice, resuspended in 1% paraformaldehyde and kept at 4°C until analyzed. Flow cytometry was performed on a FACScan (Becton Dickinson, Mountain View, CA). Fluorescence intensity was determined on at least 5,000 cells from each sample. The results were expressed as mean fluorescence intensity (MFI).
To determine the cytotoxic effect of LPC on eosinophils, propidium iodide staining was assessed in eosinophils treated with LPC. Aliquots of 0.5 × 106 eosinophils were incubated for 30 min at 37°C with 0–20 μM LPC in a total volume of 250 μl. Propidium iodide at 5 μg/ml was added to the medium of drug-treated cells, and the cell suspension was immediately analyzed by flow cytometry. Red fluorescence intensity was determined on at least 10,000 cells from each sample, and the percentage of stained cells was analyzed using the Cellquest software.
To determine the expression of TRP channels in human eosinophils by RT-PCR, mRNA enriched total RNA was isolated by a Qiagen RNeasy mini kit according to the manufacturer's instructions (Valencia, CA). Approximately 80 ng of RNA was isolated from 8 × 106 eosinophils. The entire amount of RNA obtained from each isolation was then used for cDNA synthesis. First-strand cDNA was synthesized using Bio-Rad's iScript cDNA synthesis kit (Hercules, CA). For PCR, the reaction solution (50 μl) contained 10 μl of cDNA solution, 1 μM primer (each for reverse and forward primers), 2 mM MgCl2, 0.2 mM dNTP, and 1.25 units of Taq polymerase in 1× PCR buffer supplied by Fermentas, Inc. (Hanover, MD). The cycling conditions were: 95°C for 10 min, followed by 35 cycles, each consisting of denaturation at 95°C for 1.5 min, annealing at 63°C for 2 min, and extension at 72°C for 2 min, and a final extension at 72°C for 10 min. PCR products were analyzed on 2% agarose gels and visualized by staining with ethidium bromide. Primer pairs were the same as those used by Heiner and colleagues (36). Primers for cyclophilin were used as a housekeeping gene control.
Eosinophils (106) were lysed in 40 μl of lysis buffer (50 mM Tris-HCl, 2 mM EDTA, 2 mM EGTA, 2 mM Na3VO4, 50 mM NaF, 2.5 mM DFP, 1% SDS, and protease inhibitor cocktail) and then mixed with loading buffer and boiled for 5 min. Samples then were subjected to SDS-PAGE, using 10% acrylamide gels under reducing condition (15 mA/gel). Electrotransfer of proteins from the gels to nitrocellulose membrane was achieved using a semi-dry system (400 mA, 60 min). The membrane was blocked for 60 min with 1% BSA, which then was incubated with 1:1,000 anti-TRPC6 or TRPV1 antibody diluted in Tris-buffered saline plus 0.05% Tween 20 (TBS-T) overnight. The membranes then were washed three times for 20 min with TBS-T. Anti-rabbit or mouse IgG conjugated with horseradish peroxidase was diluted 1:3,000 in TBS-T and incubated with nitrocellulose membrane for 60 min. The membrane was again washed three times with TBS-T and assayed by an enhanced chemiluminesence system (Amersham, Arlington Heights, IL).
Data are expressed as mean ± SEM. The significance of difference in adhesion between experimental groups was determined by ANOVA. When ANOVA indicated a significant difference between groups, a t test was done to determine which intergroup differences were significant. Statistical significance was claimed whenever P < 0.05.
We assessed the effect of LPC (16:0) on eosinophil adhesion to plated BSA. We have previously established that eosinophil adhesion to plated BSA is β2-integrin CD11b/CD18 dependent (5) and that BSA is a full and complete surrogate ligand for β2-integrin. LPC-induced eosinophil adhesion was concentration dependent. LPC increased eosinophil adhesion to plated BSA from 11.3 ± 1.9% for nonstimulated control to a maximum adhesion of 22.2 ± 2.1% at 4 μM (Figure 1A, P < 0.01, n = 5). Eosinophil adhesion then decreased with increasing concentrations of > 4 μM LPC. Concentrations 6 μM were toxic to the cells and caused progressive de-adhesion. At a concentration of 6 μM LPC, 81.2 ± 2.2% cells stained positive with propidium iodide as compared with control of 2.0 ± 1.5% (P < 0.01, Figure 1B, n = 4).
Because other LPC species (e.g., 18:0 and 18:1) normally are present along with 16:0 LPC in plasma and tissues, we also examined the effects of these LPC species on eosinophil adhesion. Both 18:0 and 18:1 LPC caused adhesion of eosinophils that was similar to that of 16:0 LPC, but was somewhat less potent and less efficacious for 18:1 LPC (Figure 1C, n = 5).
We next examined the effect of the charge and shape of lysophospholipids on eosinophil adhesion. Four lysophospholipids with different polar heads (ethanolamine, serine, choline, and inositol) and comparable acyl chains were used in these experiments (Figure 2A). LPI and LPS are negatively charged, whereas LPC and LPE are neutral. LPC has a large polar head with a diameter of ~ 5.2 nm and a thin hydrophobic acyl chain tail with a cross-section estimated to be 1.8–2.0 nm, and can be pictured as a cone with its base at the polar interface (37). LPC and LPI have larger head groups than LPE and LPS. As shown in Figure 2B, LPE and LPS have no effect on eosinophil adhesion. By contrast, LPI, a similar cone-shaped lipid with a large inositol head group and a single fatty acid side chain, mimicked the effect of LPC on eosinophil adhesion (toxic effect was also observed at 6μM; data not shown). These results indicate that the shape but not the charge of the lysophospholipids determines their effect on eosinophil adhesion.
We next tested whether LPC increases intracellular Ca2+ in eosinophils. LPC was applied to isolated human eosinophils in the absence of extracellular Ca2+ to study separately Ca2+ store depletion and influx. LPC was applied at t = 100 s in Ca2+-free buffer in the presence of 0.3 mM EGTA, followed by addition of 2 mM Ca2+ at t = 200 s. Unlike PAF, LPC at 4 μM did not cause Ca2+ store depletion but significantly increased Ca2+ influx in eosinophils. LPC induced sustained Ca2+ influx for at least 15 min. However, there was no propidium iodide staining even at 40 min (see below), indicating that calcium influx caused by 4 μM LPC did not result from a cytotoxic effect. A representative Ca2+ trace with 4 μM LPC is shown in Figure 3A. Typical Ca2+ store depletion transients caused by GPCR agonists, such as PAF (Figure 3B), were not detected on addition of LPC. However, Ca2+ influx was detected immediately upon addition of external calcium in LPC-treated cells at 200 s. In the absence of LPC, Ca2+ entry in eosinophils was minimal (Control, Figure 3A). By contrast, PAF applied at t = 100 s caused transient release of Ca2+ from intracellular stores followed by Ca2+ influx upon addition of external calcium at t = 200 s (Figure 3B). At 600 s LPC caused further Ca2+ influx, indicating that calcium influx caused by LPC is not mediated by store-operated calcium channels.
To elucidate the signaling cascade for LPC-induced Ca2+ entry, we next examined the involvement of G protein–coupled phospholipase C (PLC) signaling pathway. U73122, a specific inhibitor of PLC (38), was added at t = 30 s. U73122 had no effect on Ca2+ influx caused by LPC (Figure 3C). However, pretreatment with U73122 subsequently abolished Ca2+-store depletion transient caused by PAF (Figure 3D). After addition of external Ca2+, Ca2+ entry was detected in PAF-stimulated cells; LPC caused further entry from ~ 250 nM to ~ 800 nM (Figure 3D). These data indicate that G protein–coupled PLC is not the pathway that is used for Ca2+ entry caused by LPC in eosinophils.
To investigate further the mechanism of Ca2+ influx induced by LPC, we next examined the effect of lanthanum and SKF 96365, a store-operated Ca2+ channel blocker (45) in human eosinophils. Pretreatment eosinophils with 100 μM La3+ (Figure 3E), a nonselective TRP channel blocker (39), attenuated the rate of Ca2+ influx caused by LPC. By contrast, SKF96365, had no effect on calcium influx caused by LPC (Figure 3F). 2-APB, another store-operated calcium channel blocker, did not block Ca2+ entry either (data not shown).
We next examined the effect of the shape of LPC on calcium influx in eosinophils. Previous study has shown that externally applied LPC to lipid bilayers increases large mechanosensitive channel (MscL) activity (40). PE (which has a small ethanolamine head group and two acyl chains as a large tail group, Figure 3G, inset), a lipid with shape oppositely oriented to that of LPC, neutralizes the effect of LPC on MscL activity (40). In our study, preincubation of eosinophils with 10 μM PE blocked almost completely Ca2+ entry caused by LPC (Figure 3G), indicating that the effect of LPC on eosinophils also is caused by the same mechanism of outer membrane insertion of LPC.
Finally, we investigated whether BSA neutralized LPC-induced Ca2+ entry. LPC in plasma is present mainly in albumin- and lipoprotein-bound form (41). Pretreatment of eosinophils with 3 μM BSA abolished effect of LPC on Ca2+ entry (Figure 3F). These data suggest that free LPC, but not the protein bound LPC, has the ability to cause Ca2+ entry.
Since PE blocked Ca2+ entry caused by LPC, we then examined the effect of PE on eosinophil adhesion caused by LPC. 18:1 dioleoyl-phosphatidylethanolamine blocked eosinophil adhesion caused by LPC (Figure 4A). Adhesion of nonstimulated eosinophils to plated BSA was 10.4 ± 2.1%. Adhesion caused by 4 μM LPC was blocked by PE from 23.2 ± 4.0 to 11.6 ± 1.1% at 3 μM, and was further blocked to 9.5 ± 1.5 at 10 μM (P < 0.01 versus control).
Previous investigations have shown that albumin binds to lysophospholipids, thereby sequestering these lipids away from the cell surface (41, 42). We therefore examined the effect of increasing concentrations of soluble albumin on eosinophil adhesion caused by LPC. At concentrations of BSA approaching 1–3 μM, eosinophil adhesion caused by LPC was completely blocked (Figure 4B).
To examine whether the presence of extracellular Ca2+ was required for LPC-induced eosinphil adhesion, eosinophil adhesion in response to LPC was measured in the calcium-free HBSS +0.1% gelatin in the presence of 0.3 mM EGTA (Figure 4C). In the regular HBSS buffer, which contains 1.3 mM Ca2+, LPC at 4 μM induced eosinophil adhesion from buffer control of 8.2 ± 2.0% to 25.1 ± 2.0% (P < 0.01). In the absence of extracellular Ca2+, no adhesion was induced by LPC (Figure 5C, P = NS versus control without calcium).
We next examined the role of two previously postulated mechanisms of integrin-mediated adhesion: (1) up-regulation of surface integrin molecules, and (2) the affinity/conformational changes of the β2-integrin (43). We first determined the effect of LPC in regulating Mac-1 (CD11b/CD18) expression by flow cytometry. Stimulation of eosinophils with LPC did not increase significantly the expression of the CD11b α-chain of Mac-1 on the eosinophil surface (baseline control of 18.0 ± 1.3 versus 19.5 ± 2.1 MFI for treatment [P = NS]) (Figures 5A and 5E). Treatment of eosinophils with 10 μM PE had no effect on CD11b surface expression either (Figure 5E, 17.8 ± 1.9 versus nonstimulated control, P = NS). By contrast, CD11b expression was up-regulated by PAF, from 18.0 ± 1.3 to 31.6 ± 2.0 (P < 0.01). Pretreatment of PE did not prevent the surface up-regulation of CD11b caused by PAF (Figures 5C and 5E).
We next determined whether LPC up-regulates the change to active conformation for Mac-1 (CD11b/CD18). The activated conformation of Mac-1, as identified by the mAb CBRM1/5 (2), was induced after treatment with LPC from 19.3 ± 1.3 to 32.5 ± 1.2 MFI (P < 0.01, Figure 5B). PE alone had no effect on the expression of active CD11b (Figure 5F); however, it blocked the active conformation of Mac-1 caused by LPC, from 32.5 ± 1.2 to 20.3 ± 1.4 MFI (Figures 5B and 5F, P < 0.01). By contrast, PAF-mediated up-regulation of active conformational change of Mac-1 (Figures 5D and 5F) was not attenuated by PE.
As our data suggested the involvement of non–store-operated Ca2+ channels, we next investigated the potential of TRP channel involvement is this phenomenon. Recent studies have suggested that LPC may incorporate into the outer membrane of the lipid bilayer, thereby changing the membrane curvature and activating the mechanosensitive ion channel within the lipid membrane (30, 40). We examined the profile of TRP channel expression in human eosinophils. Specific primer pairs were used to screen for the presence of TRP mRNA species in the highly purified human eosinophils. The expression profile for members of the TRP family is illustrated in a representative gel of RT-PCR products shown in Figure 6A. PCR products for TRPC6 and TRPV1 were found in all eosinophil samples. To confirm TRPC6 and TRPV1 protein expression, we examined eosinophil lysates by Western blot analysis. A specific antibody for TRPC6 revealed a strong band of ~ 105 kD (Figure 6B). Antibody for TRPV1 revealed a band of ~ 110 kD in NIH3T3 cells (Figure 6C, lanes 2 and 3); however, only very weak expression of TRPV1 was ever found in human eosinophils (Figure 6B, lane 2, and Figure 6C, lane 1). To determine if this very weak band was either nonspecific or nonfunctional, we administered 100 μM capsaicin, a known activator of TRPV1, to aliquots of 106 eosinophils for up to 10 min. No calcium influx was demonstrated.
The major findings in this study are as follows. (1) Extracellular LPC elicits increased β2-integrin–mediated eosinophil adhesion to immobilized BSA. This effect is mimicked by the same cone-shaped lipid, LPI, and cancelled by PE, a lipid with shape opposite to that of LPC. (2) In contrast to G protein receptor agonists, LPC increases Ca2+ entry without having any detectable effect on Ca2+ store depletion. (3) This Ca2+ influx is blocked by preincubation with either PE or La3+, a general blocker of mechanosensitive channels. (4) LPC causes active conformational change of CD11b that is prevented by preincubation with PE. These findings provide the evidence that LPC causes a non–store-operated Ca2+ influx, which causes activation of β2-integrin and subsequent eosinophil adhesion.
The mechanism by which LPC alters cell function has remained elusive. Our studies indicate that rather than GPCR activation as suggested previously (18, 19, 44), Ca2+ influx through a non–store-operated calcium channel is important for the induction of eosinophil adhesion by LPC. We show that eosinophil adhesion caused by LPC is regulated by Ca2+ influx. The inhibition of Ca2+ influx by PE (Figure 3G), BSA (Figure 3H), or in the absence of extracellular Ca2+ (Figure 3) corresponded to the decreased eosinophil adhesion (Figure 4) caused by these agents. Other evidence suggests that Ca2+ influx caused by LPC is independent of GPCR. (1) The marked Ca2+ entry (Figure 3A) in the absence of store depletion in response to LPC suggests a mechanism other than G protein receptor–dependent store depletion and subsequent store-operated calcium ion entry, such as in the case of fMLP or PAF. (2) LPC caused further Ca2+ entry in the presence of PAF, which caused store-operated calcium entry, suggesting that LPC-induced Ca2+ influx comes through routes other than store-operated calcium channels (Figure 3B). (3) Pharmacologic inhibition of PLC by U73122 had no effect on calcium entry caused by LPC. This excludes the role of Gi or Gq-PLC pathway in LPC-induced eosinophil adhesion. Gs-mediated cAMP production has been linked to decreased eosinophil adhesion, excluding the possible involvement of Gs in LPC-induced eosinophil adhesion (45). Involvement of Gα13 is also unlikely, as Gα13 caused-Rho activation promotes eosinophil detachment (46). (4) The calcium entry caused by LPC was blocked by La3+, suggests a role of a mechanosensitive cation channel. (5) LPC induced eosinophil adhesion in a narrow concentration ranging from 1–4 μM, compared with receptor-mediated agonist, which activates eosinophils over a range of 2–3 log concentrations (5, 47, 48). The narrow dose–response curve of LPC on eosinophil adhesion also argues against the likehood of its action on a cell surface receptor. (6) Prior reports on the putative receptor for LPC, including G2A and GPR4, have been retracted from publication (27, 28). Altogether, our data suggest that Ca2+ entry and subsequent eosinophil adhesion caused by LPC is mediated by an integral membrane cation channel rather than by a GPCR.
Perozo and coworkers have hypothesized a mechanism by which LPC causes activation of mechanosensitive channels (40). They propose that asymmetry in the lateral pressure between the two leaflets of the bilayer initiates the sequence of mechanical transduction steps that leads to the open state of mechanosensitive channels. According to the authors' hypothesis, the cone-shaped LPC causes the release of intrabilayer lateral pressure due to its insertion into the outer monolayer of the plasma membrane. By sensing this pressure change, mechanosensitive channels undergo conformational changes and subsequently open. Application of Perozo's hypothesis leads us to the conclusion that eosinophil calcium entry channels are activated by the same mechanism. Our data support this hypothesis. We find that LPI, a lipid with shape similar to LPC, mimics the effect of LPC on eosinophil adhesion (Figure 2). PE, a lipid with a shape opposite to that of LPC, blocked the effect of LPC on Ca2+ entry (Figure 3), eosinophil adhesion (Figure 4), and CD11b active conformational change (Figure 5). As LPC activates mechanosensitive channels on both prokaryotic and eukaryotic cells, we believe that LPC may represent an evolutionarily conserved crude modulator of integral membrane proteins.
We find that preincubation of eosinophils with soluble BSA substantially attenuated the bioactivity of LPC. Soluble albumin has been shown to contain one to three high-affinity binding sites for LPC (49). LPC in plasma is present mainly in albumin- and lipoprotein-bound form (41). These forms may play an important role in delivering fatty acids and choline to tissues such as the brain (41), but may be in a form that is unable to stimulate eosinophil functions. It has been shown that some of the effects of LPC are decreased in the presence of albumin (50). Thus, the functionally available concentration of LPC in vivo, and the stimulatory action of eosinophil function, may be controlled by the low concentrations of free LPC. Our results shown in Figure 4B appear to support this notion. The presence of 3 μM BSA also reversed the ability of LPC to cause calcium influx (Figure 2H). The physiologically relevant concentrations of LPC in vivo will be better understood when estimates of unbound LPC concentrations in specific tissues can be reliably made. These results suggest that effective proinflammatory activity of LPC can occur in tissues either in which there is elevated LPC concentration that exceeds the binding capacity of albumin or under conditions of decreased albumin levels. We do not think immobilized BSA on 96-well plates interfered with the effect of LPC on eosinophil adhesion, as free LPC was preincubated with eosinophils first.
Compared with Ca2+ release from intracellular stores, Ca2+ entry still is not well elucidated. Members of the TRP family of ion channels represent the first potential candidates for calcium entry channels in nonexcitable cells. In this study, we found that TRPC6 and TRPV1 both are expressed as mRNA in eosinophils. These two mRNAs are also found in human neutrophils (36). We further confirmed the protein expression of TRPC6 by Western blot. By contrast, we found weak or no TRPV1 protein expression in eosinophils. Capsaicin, a TRPV1 agonist, also caused no increase in intracellular calcium in eosinophils. Although the role of TRPC6 could not be defined specifically, as selective blockade is not yet technically possible, a role for TRPV1 as a regulator of Ca2+ influx in these studies was excluded.
We conclude that the naturally occurring lysophospholipid LPC causes a non–store-operated Ca2+ influx, which subsequently activates CD11b/CD18 to cause eosinophil adhesion. This effect of LPC on eosinophil adhesion appears to be independent of GPCR. This newly identified LPC signaling mechanism could provide insight into the molecular mechanisms of the multiple actions of LPC in cells involved in various inflammatory diseases. However, the role of LPC blockade as a potential therapeutic approach for attenuating eosinophilic inflammation or inflammation in other inflammatory cells remains to be elucidated.
This work was supported by National Institute of Allergy and Infectious Disease grant AI52109 (X.Z.), National Heart, Lung, and Blood Institute (NHLBI) grant HL-46368, by NHLBI SCOR grant HL-56399 (A.R.L), and by a grant from the GlaxoSmithKline Center of Excellence.
Originally Published in Press as DOI: 10.1165/rcmb.2006-0391OC on January 11, 2007
Conflict of Interest Statement: X.Z. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. J.L. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. S.B. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. L.Z. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. P.V.U. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. V.N. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. N.M.M. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. A.R.L. has received $3,000 in the past 3 years for serving on a GlaxoSmithKline Advisory Board, $13,000 per year for a Merck Advisory Board, $24,000 per year for Broncus/Asthmatx Advisory Board, and $3,500 for a Pfizer Advisory Board.