The P2X7 receptor (P2X7R), an ATP-gated ion channel, plays essential roles in the release and maturation of IL-1β in microglial cells in the brain. Previously, we found that lysophosphatidylcholine (LPC) potentiated P2X7R-mediated intracellular signals in microglial cells. In this study, we determined whether the lysophospholipids, i.e., LPC and sphingosylphosphorylcholine (SPC), modulate the ATP-induced release and processing of IL-1β mediated by P2X7R in mouse MG6 microglial cells. LPC or SPC alone induced the release of precursor (pro-IL-1β) and mature IL-1β (mIL-1β) from LPS-primed MG6 cells, possibly due to lytic functions. However, these lysophospholipids inhibited ATP-induced caspase-1 activation that is usually followed by the release of mIL-1β. Conversely, ATP inhibited the release of pro-IL-1β and mIL-1β induced by LPC/SPC. This suggests that lysophospholipids and ATP mutually suppressed each function to release IL-1β. P2X7R activation resulted in microtubule reorganization in the MG6 cells that was blocked in the presence of LPC and SPC. LPC/SPC reduced the amount of activated RhoA after stimulation with ATP, implying that these lysophospholipids block ATP-induced microtubule reorganization by interfering with RhoA activation. In addition, the microtubule inhibitor colchicine inhibited ATP-induced release of mIL-1β similar to that of LPC and SPC. This suggests that the impairment of the microtubule reassembly may be associated with the inhibitory effects of LPC/SPC on ATP-induced mIL-1β release. Mutual suppression by ATP and LPC/SPC on the maturation of IL-1β was observed in LPS-primed primary microglia. Collectively, these data suggest opposing functions by lysophospholipids, either proinflammatory or anti-inflammatory, in regard to the maturation and release of IL-1β from microglial cells.

Microglia are immunocompetent cells in the CNS and share many of the functional properties of macrophages in peripheral tissues (1). In the normal adult brain, microglia are in a resting state and monitor the brain microenvironments using their motile processes (2). If neuronal injury or infections by microorganisms occurs, resting microglia transform into an activated “amoeboid” shape and secrete proteases, neurotrophic factors, and various cytokines (3). Microglia physiologically play protective roles against neuronal damage, whereas in some pathological conditions the overactivated microglia produce various neurotoxic substances and promote neuronal cell death. The increased levels of inflammatory cytokines, e.g., IL-1β, IL-6, and TNF-α, secreted from microglia in brains likely correlates with the progression of neurodegenerative disorders such as Alzheimer’s disease and Parkinson’s disease (4, 5).

IL-1β is a potent proinflammatory cytokine primarily produced in the brain microglia. The secretion of IL-1β from these cells is stringently controlled because it is a primary mediator for inflammation (6, 7). IL-1β is synthesized as a biologically inactive procytokine (pro-IL-1β; ∼33 kDa) that accumulates in the cell cytoplasm. Pro-IL-1β is then processed by activated caspase-1, the primary protease responsible for intracellular IL-1β processing into the biologically active mature form (mIL-1β;3 ∼17 kDa) that is released into the extracellular space. Caspase-1 is also synthesized as an inactive 45-kDa procaspase-1 that is made active by cleavage of the C terminus into p10 and p20 fragments to form a heterotetramer (two p10 and two p20 subunits) (8). Bacterial endotoxin, LPS, is commonly used to induce gene expression and protein synthesis for pro-IL-1β in microglial cells (9); however, this stimulation is inefficient to trigger a rapid release of mIL-1β. Pro-IL-1β lacks the signal sequence for transport using the classical exocytotic process (10) where the maturation and release of IL-1β are considered to be modulated by another mechanism (11).

ATP induces the rapid release of mIL-1β from microglia at relatively higher concentrations (>1 mM) (9). The P2X7 receptor (P2X7R), one of the purinergic receptor families highly expressed in microglia, responds to this higher ATP concentration and facilitates the activation of caspase-1, and this is followed by the processing and release of IL-1β in the LPS-primed microglial cells (9). The efflux of K+ from the cells induced by P2X7R activation is thought to be essential for the activation of caspase-1 as well as the release of mIL-1β in many kinds of cells, including monocytes and macrophages (9, 12, 13). Although P2X7R-dependent mIL-1β release requires other intracellular signals, i.e., Ca2+ influx (14) and the activation of phospholipase C and A2 (12), the relevance of these signals may vary among the cell types.

Several models are proposed to explain nonclassical secretion of mIL-1β using the P2X7R-dependent pathway from cells of the monocyte/macrophage lineage (11). The simple and oldest model is that necrotic cell death and lysis induced by stimuli triggers the processing and release of IL-1β. However, several recent studies show that the pathways leading to mIL-1β release are not explained by simple cell lysis; but are more sophisticated. Therefore, a second model was proposed where pro-IL-1β and procaspase-1 are transported into lysosomal vesicles and processed, and then the vesicles containing the mIL-1β are released from the cell using exocytosis (12). In the third model, P2X7R stimulation initiates the local accumulation of procaspase-1 and pro-IL-1β within the microdomain of the subplasma membrane cytosol and then the microvesicles containing the mIL-1β are formed and rapidly shed from the cell surface (15, 16). Most recently, two alternative models were proposed for macrophages: 1) ATP-induced pro-IL-1β processing occurs in the cytosol and mIL-1β may directly pass through the plasma membrane using molecularly undefined transporters (17) and 2) the formation of endosome-derived recycled multivesicular bodies may contain exosomes entrapping mIL-1β to extracellularly release of mIL-1β (18). Although microvesicle shedding may explain the larger part of the mechanism for ATP-induced mIL-1β release in microglial cells (19), the modulatory mechanisms for the processing and release of IL-1β via P2X7R activation remain to be determined.

We recently reported lysophosphatidylcholine (LPC) potentiates the P2X7R-mediated signals, i.e., Ca2+ influx, membrane pore formation, and p44/42 MAPK activation in microglial cells (20). Additionally, LPC is shown to activate nonselective cation channels and to induce the K+ efflux through the activation of Ca2+-dependent K+ channels, and this ultimately leads to the release of mIL-1β in microglia (21). Similarly, sphingosylphosphorylcholine (SPC), another bioactive lysophospholipid, as well as LPC are reported to enhance P2X7R-mediated IL-1β release in THP-1 cells by increasing the binding affinity of agonists to P2X7R (22). These studies demonstrate the proinflammatory features of these lysophospholipids in immune cells. In contrast, LPC or SPC reduces the release of proinflammatory cytokines, i.e., IL-1β and TNF-α, in experimental sepsis in vivo to protect mice against lethality (23, 24, 25). This suggests its possible role as an anti-inflammatory factor.

To determine the roles of lysophospholipids in microglial activation, here we investigate the effects of two lysophospholipids, LPC and SPC, on ATP-induced IL-1β release from immortalized microglial MG6 cells and primary mouse microglia. We demonstrate that LPC and SPC inhibit the P2X7R-dependent maturation of IL-1β in these microglial cells possibly using RhoA-dependent mechanisms. Our data suggest that LPC or SPC has essential physiological roles in the brain not only as a proinflammatory agent, but also as anti-inflammatory factors by suppressing the P2X7R-dependent maturation and release of IL-1β in microglia.

ATP, oxidized ATP (oATP), LPC (1-palmitoyl-sn-glycero-3-phosphocholine), SPC, LPS, nigericin, PD98059, A23187, colchicine, anti-β-tubulin Ab, and BSA were purchased from Sigma-Aldrich. LPC and SPC were dissolved in 50% ethanol at 25 mM and stored in aliquots at –30°C. HRP-conjugated goat anti-rabbit and anti-mouse Ig Abs were purchased from Valeant Pharmaceuticals. Monoclonal anti-mouse IL-1β Ab (MAB401), biotinylated anti-mouse IL-1β Ab (BAF401), and recombinant mouse mIL-1β were purchased from R&D Systems. HRP-streptavidin conjugate was obtained from Zymed Laboratories. Anti-caspase-1 and anti-cathepsin D polyclonal Abs were purchased from Santa Cruz Biotechnology. Anti-actin Ab was purchased from Chemicon International. ECL Plus kit and x-ray films were purchased from Amersham Pharmacia Biotech. Potassium-binding benzofuran isophtalate acetomethylester was purchased from Invitrogen. Caspase-1 Inhibitor II (Ac-YVAD-CMK), Y27632, and HA1077/fasudil were purchased from Merck-Biosciences.

A c-myc-immortalized mouse microglial cell line, MG6, was established from a primary cultured microglia as described in our previous study (26). The MG6 cells were routinely maintained in growth medium composed of DMEM containing 10% FBS supplemented with 100 μM 2-ME, 10 μg/ml insulin, 100 μg/ml streptomycin, and 100 U/ml penicillin in 100-mm petri dishes (BD Biosciences).

Primary microglial cells were obtained from cultured brain cells from neonatal C57BL/6 mice (CLEA Japan). Mixed brain cell cultures were prepared in tissue culture flasks according to a protocol described previously (27). After 10–20 days of culture, microglia were harvested by shaking the culture flasks for 1.5 h at 80 rpm. Harvested cells were seeded in 100-mm petri dishes and incubated for 10 min at 37°C. After removing nonadherent cells using a single wash with PBS, the attached microglia were maintained in the growth medium. The next day, the microglial cells were removed using PBS containing 1 mM EDTA and seeded in 24-well plates (BD Biosciences) for the IL-1β release assay or caspase-1 activation determination.

MG6 cells were seeded in 24-well plates at a density of 3 × 105 cells/well. The next day, the cells were incubated with growth medium containing LPS (1 μg/ml) for 4 h, washed with PBS, and then the growth medium was replaced with 500 μl of HEPES-buffered salt solution (HBSS: 145 mM NaCl, 2.5 mM KCl, 1 mM MgCl2, 1.8 mM CaCl2, 20 mM HEPES, 10 mM glucose, 0.01% BSA, pH 7.4) containing the test reagents. After 30 min of incubation at 37°C, the culture supernatant was collected into 1.5-ml tubes and stored at −30°C until used. After brief centrifugation to remove the debris, 100-μl aliquots of culture supernatant samples were assayed for IL-1β using a sandwich ELISA according to the manufacturer’s instructions (R&D Systems). A 96-well plate was coated with 2 μg/ml monoclonal anti-mouse IL-1β Ab (MAB401) at 4°C overnight and then blocked with 1% BSA in PBS for 1 h. The plates were washed three times with PBS containing 0.2% Tween 20 (PBST). Aliquots of supernatant samples or recombinant mouse IL-1β standards were diluted to 100 μl with HBSS, added to the blocked wells, and incubated for 2 h at room temperature. The plates were washed three times with PBS and 100-μl aliquots of 0.1 μg/ml biotinylated mouse IL-1β affinity-purified polyclonal detection Ab (BAF401) were added and incubated for 2 h. After a further three washes with PBST, the captured immune complexes were colorimetrically detected using subsequent incubations with HRP-streptavidin conjugate and SureBlue TMB 1 component microwell peroxidase substrate (Kirkegaard & Perry Laboratories) to detect the biotinylated Ab. Finally, stop solution (1 M H2SO4) was added and the absorbance at 450 nm in each well was measured using a microplate reader (VersaMax; Molecular Devices). A standard curve was created using serial dilutions of recombinant mouse IL-1β and used to calculate the assay results. The experiments were independently performed six times and the data are expressed as the mean value ± SEM.

MG6 cells and primary microglia were seeded into 24-well plates at a density of 3 × 105 cells/well. The next day, the cells were incubated with the growth medium containing LPS (1 μg/ml) for 4 h, washed with PBS, and then the growth medium was replaced using either 500 μl (for IL-1β release assay) or 250 μl (for caspase-1 activation assay) of HBSS containing the test reagents. To determine the effect of oATP, the cells were incubated with LPS-containing growth medium in the presence or absence of 300 μM oATP. Caspase-1 inhibitor was added into the HBSS combined with stimulating reagents to assess its inhibitory effect. After 30 min of incubation at 37°C, the culture supernatant was collected into 1.5-ml tubes and stored at −30°C until used. To prepare the cell extraction, the cells were lysed with 400 μl of radioimmunoprecipitation assay (RIPA) buffer (50 mM Tris-HCl (pH 7.5), 150 mM NaCl, 0.5% Triton X-100, 0.5% sodium deoxycholate, containing a complete mini protease inhibitor tablet (Roche)) to detect IL-1β release and 250 μl of RIPA buffer to detect caspase-1 activation. The cell lysates were stored at −30°C until used. After the cell debris was removed by centrifugation, equal volumes of culture supernatant (16 μl to detect IL-1β and caspase-1) and cell lysate (4 μl for IL-1β and 16 μl for caspase-1) were separated by SDS-PAGE (15%) and electroblotted onto a polyvinylidene difluoride (PVDF) membrane (Immobilon-P; Millipore). To detect the activated caspase-1 in the culture supernatant of MG6 cells, the proteins in the 100-μl supernatant were concentrated using centrifugation after the addition of four volumes of methanol and then they were applied to SDS-PAGE. The membranes were blocked with 5% nonfat milk in TBST for 1 h. To detect the processing of IL-1β, the membranes were incubated with biotinylated anti-IL-1β Ab (1/1000) for 1 h followed by incubation with HRP-streptavidin conjugate (1/5000) for 20 min. To detect caspase-1 activation, membranes were incubated with anti-caspase-1 Ab (1/1,000) in TBST for 1 h followed by incubation with HRP-conjugated goat anti-rabbit Ab (1/10,000) for 1 h. The target proteins were revealed using an ECL Plus kit and detected on x-ray film.

MG6 cells were seeded in 24-well plates at a density of 3 × 105 cells/well. The next day, the cells were incubated with growth medium containing LPS (1 μg/ml) for 4 h, washed with PBS, and then the growth medium was replaced using 500 μl of HBSS containing the test reagents. After 30 min of incubation at 37°C, the culture supernatant was collected into 1.5-ml tubes. To prepare the cell extract, the cells were lysed in 500 μl of HBSS containing 1% Triton X-100. The samples were used for the measurement of LDH activity without freezing. Either 50 μl of culture supernatant or 2 μl of cell lysate diluted to 50 μl with HBSS was analyzed for LDH activity using the LDH Cytotoxicity Detection Kit (Takara Bio). The data are expressed as the percentage of the total cytosolic LDH obtained from nontreated MG6 cell lysates.

Once harvested, MG6 cells were washed and resuspended in HBSS. [K+]i was measured using PBFI fluorescence as described previously (28). In brief, potassium-binding benzofuran isophtalate acetomethylester was added to the cell suspension (10 μM) and the cells were loaded for 1 h at 37°C. After two washes with HBSS, 1.5 × 106 cells/ml were used in each set of experiments and monitored using a CAF-110 spectrofluorometer (Japan Spectroscopy) at 37°C with continuous stirring at 400 rpm. The PBFI-loaded cells were excited at 340 and 380 nm and the fluorescence was measured at 500 nm. The fluorescence ratio of F340:F380 was used to estimate the change in [K+]i. Each [K+]i measurement was repeated at least three times.

MG6 cells (1 × 106 cells) were cultured in 35-mm dishes for 24 h and further incubated with growth medium containing LPS (1 μg/ml) for 4 h. Then, the subcellular extracts were prepared from the cell monolayer using a ProteoExtract Subcellular Proteome Extraction Kit (Calbiochem) according to the manufacturer’s instructions. Four fractions containing specific proteins localized within cytosol (F1), membrane/organelle (F2), nucleus (F3), and cytoskeleton (F4) were obtained. Each fraction (16 μl) was resolved using 15% SDS-PAGE and electroblotted onto a PVDF membrane for immunoblotting with anti-IL-1β and anti-cathepsin D Abs.

MG6 cells were seeded at a density of 1 × 105 cells per well in 8-well chamber slides (BD Biosciences). The next day, the cells were incubated with growth medium containing LPS (1 μg/ml) for 4 h, washed with PBS, and the medium was replaced with HBSS (250 μl) containing the test reagents. To determine the effect of oATP, the cells were incubated with LPS-containing growth medium in the presence or absence of 300 μM oATP. After 10 min of incubation at 37°C, the cells were fixed using a 250-μl formalin/PBS solution (3.7% formalin) for 15 min at room temperature. After fixation, cells were treated with 1% Triton X-100 for 10 min followed by incubation with 2% hydrogen peroxide for 5 min at room temperature to block endogenous peroxidase activity. After washing with PBST, the slides were blocked with 5% normal goat serum and 1% BSA (fraction V) in PBS for 30 min and then incubated with either anti-β-tubulin (1/200) or anti-actin (1/400) Ab for 1 h at room temperature. After rinsing the slides with PBST, the EnVision system (DakoCytomation) was used that contains a polymeric conjugate consisting of a large number of secondary Abs (goat anti-mouse and goat anti-rabbit) bound directly to a dextran backbone containing HRP. After further rinsing the slides with PBST, the Ab-Ag reaction was visualized with 3,3′-diaminobendizine tetrahydrochloride as the chromogen. After an additional wash with distilled water, the slides were dehydrated through an ethanol series and mounted with a coverglass using Mount-Quick (Daido Sangyo). The immunostained slides were observed using a Leica microscope.

MG6 cells were seeded at a density of 4 × 105 cells/well in 24-well plates. The next day, the cells were incubated with growth medium containing LPS (1 μg/ml) for 4 h, washed with PBS, and the medium was replaced using HBSS (500 μl) containing the test reagents. Additionally, the cells were cultured in serum-free DMEM for 16 h and stimulated using HBSS containing the test reagents. After 30 min of incubation at 37°C, RhoA activation was measured using a Rho Activation Assay Bio Kit (Cytoskeleton) according to the manufacturer’s instructions, with minor modifications. In brief, the cells were lysed with 150 μl cell lysis buffer; the lysates were cleared by centrifugation at 14,000 × g for 5 min at 4°C and100 μl of lysate was transferred to a new tube. Rhotekin beads (25 μg) were added to the supernatant followed by 1 h of incubation at 4°C with a rotator to bind all activated RhoA to the beads. The beads were then washed with wash buffer and incubated with 5× SDS sample buffer at 95°C to dissociate the activated RhoA from the rhotekin beads. The sample was separated on 15% SDS-PAGE followed by immunoblotting for RhoA.

Quantitative analysis of band intensities for immunoblot was performed using the image processing software Image J 1.38 version for the Macintosh (National Institutes of Health). The values are shown as means ± SEM. The mean values were analyzed with the Student unpaired t test or a one-way ANOVA and followed using the Scheffe’s F post hoc test with the Statview II statistical package for the Macintosh (Abacus Concepts). Statistical significance was set at p < 0.05.

We initially investigated the effects of LPC on ATP-induced IL-1β release from MG6 cells using ELISA. After stimulation with 2 mM ATP, the amount of IL-1β released into the culture supernatant increased in LPS-primed MG6 cells (Fig. 1). Priming with LPS was required to induce ATP-dependent IL-1β release in these cells as previously reported in other microglial cell lines (9). Thus, all through this study, MG6 cells or primary microglia were primed with LPS to stimulate the release of IL-1β. The IL-1β release induced by ATP was inhibited by a potent P2X7R antagonist, oATP (data not shown), showing P2X7R is involved in ATP-induced IL-1β release from MG6 cells. Administering LPC alone induced IL-1β release from LPS-primed MG6 cells at 60–100 μM, although the stimulatory potency of LPC was weaker than using ATP (Fig. 1,A). Because our previous report demonstrated LPC potentiates ATP-induced intracellular signaling mediated by P2X7R in MG6 cells (20), we expected LPC would also promote the ATP-induced release of IL-1β in these cells. However, the application of LPC dose-dependently suppressed the release of IL-1β induced by 2 mM ATP (Fig. 1,A). In addition, 1 mM ATP significantly inhibited the release of IL-1β from MG6 cells induced by 80 and 100 μM LPC (Fig. 1 A). Therefore, LPC and ATP mutually suppress IL-1β release from MG6 cells.

FIGURE 1.

LPC and SPC inhibit ATP-induced IL-1β release in LPS-primed MG6 cells. MG6 cells (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were stimulated for 30 min using HBSS containing ATP and LPC (A) or SPC (B) at the concentrations indicated. The culture supernatant was collected and analyzed using ELISA to determine the amount of IL-1β. The amount of IL-1β released in the supernatant was expressed as the mean value ± SEM (n = 6). ∗, p < 0.05 and ∗∗, p < 0.01.

FIGURE 1.

LPC and SPC inhibit ATP-induced IL-1β release in LPS-primed MG6 cells. MG6 cells (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were stimulated for 30 min using HBSS containing ATP and LPC (A) or SPC (B) at the concentrations indicated. The culture supernatant was collected and analyzed using ELISA to determine the amount of IL-1β. The amount of IL-1β released in the supernatant was expressed as the mean value ± SEM (n = 6). ∗, p < 0.05 and ∗∗, p < 0.01.

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We then investigated the effects of SPC, a structurally similar lysophospholipid to LPC, on ATP-induced release of IL-1β from LPS-primed MG6 cells. SPC alone induced IL-1β release in a dose-dependent manner at 60–100 μM (Fig. 1,B). SPC suppressed the release of IL-1β induced by 2 mM ATP and its inhibitory effect was more potent than using LPC (Fig. 1,B). As observed with LPC, the IL-1β release induced by SPC was inhibited by 1 mM ATP (Fig. 1 B), suggesting the mutual suppression of IL-1β release by these two ligands in MG6 cells. The data demonstrate that both LPC and SPC exhibit inhibitory effects on the IL-1β release induced by ATP and, conversely, ATP has the ability to suppress the LPC/SPC-induced IL-1β release in MG6 cells.

To distinguish the two forms of IL-1β, the 33-kDa pro-IL-1β and the proteolytically processed 17-kDa mIL-1β known to be released from the cells (9), we analyzed IL-1β released from LPS-primed MG6 cells and primary microglia using immunoblot. LPC alone dose-dependently induced the release of large amounts of pro-IL-1β as well as a small amount of mIL-1β into the culture supernatant (Fig. 2,A). This indicates both precursor and mature forms of IL-1β were released from microglial cells by LPC. In primary microglia, LPC induced significant production of mIL-1β at lower concentrations (∼20 μM) than those observed in the MG6 cells (Fig. 2,A). The LPC-induced release of mIL-1β was reduced when simultaneously treated with 1 mM ATP in MG6 cells (Fig. 2,A). This is in agreement with the data obtained using ELISA (Fig. 1,A). ATP triggered the release of both pro- and mIL-1β at 2 mM (Fig. 2,A). LPC dose-dependently suppressed the release of mIL-1β induced by ATP in MG6 cells and primary microglia, whereas the release of a novel 20-kDa form of IL-1β was increased in a dose-dependent manner with the lipid in primary cells (Fig. 2 A).

FIGURE 2.

LPC and SPC inhibit ATP-induced mIL-1β release and caspase-1 activation in LPS-primed microglial cells. MG6 cells and primary microglia (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were stimulated for 30 min using HBSS containing ATP and LPC or SPC at the concentrations indicated. The culture supernatant was collected and the cells were lysed with RIPA buffer. Equal volumes of supernatant (sup) or cell lysates were loaded into each lane for SDS-PAGE. After transfer to a PVDF membrane, the blots were probed with anti-IL-1β (A, D, and E) or anti-caspase-1 (C) Ab. LPC and SPC primarily induced the release of pro-IL-1β (pro) and procaspase-1 (p45) in MG6 cells and primary microglia (primary) (A and C). Because LPC or SPC alone induced the release of mIL-1β weakly in MG6 cells (A), the production of activated caspase-1 p10 (p10) was not detected using this experimental condition (C). LPC induced the significant release of mIL-1β (A) and activated caspase-1 p10 (C) in primary cells. In the primary microglia, the 20-kDa form of IL-1β (20k) appeared using the cotreatment with ATP and LPC/SPC (A and B). LPC/SPC inhibited ATP-induced mIL-1β release (A) and production of the p10 fragment (C) in a dose-dependent manner. If caspase-1 inhibitor (CaspI) was added to HBSS, the ATP-induced release of mIL-1β was inhibited showing the involvement of this protease in the maturation of IL-1β in MG6 cells (D). Treatment of the cells with 300 μM oATP during the LPS priming abolished the ATP-induced release of mIL-1β, suggesting the involvement of P2X7R, whereas the P2X7R function was not involved in LPC-induced release of pro- and mIL-1β (E). The asterisks in C indicate nonspecific bands corresponding to BSA contained in HBSS. The immunoblots are representative of at least three independent experiments. B, band intensities corresponding to mIL-1β (▪) and the 20-kDa form (□) in the supernatant from primary microglia were quantified in three independent experiments, and the data are indicated as a percentage of the amount of mIL-1β or the 20-kDa form obtained from the culture supernatant after the stimulation using 2 mM ATP in each set of experiments. The data are expressed as the mean value ± SEM. ∗, p < 0.05; ∗∗, p < 0.01; and ∗∗∗, p < 0.001.

FIGURE 2.

LPC and SPC inhibit ATP-induced mIL-1β release and caspase-1 activation in LPS-primed microglial cells. MG6 cells and primary microglia (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were stimulated for 30 min using HBSS containing ATP and LPC or SPC at the concentrations indicated. The culture supernatant was collected and the cells were lysed with RIPA buffer. Equal volumes of supernatant (sup) or cell lysates were loaded into each lane for SDS-PAGE. After transfer to a PVDF membrane, the blots were probed with anti-IL-1β (A, D, and E) or anti-caspase-1 (C) Ab. LPC and SPC primarily induced the release of pro-IL-1β (pro) and procaspase-1 (p45) in MG6 cells and primary microglia (primary) (A and C). Because LPC or SPC alone induced the release of mIL-1β weakly in MG6 cells (A), the production of activated caspase-1 p10 (p10) was not detected using this experimental condition (C). LPC induced the significant release of mIL-1β (A) and activated caspase-1 p10 (C) in primary cells. In the primary microglia, the 20-kDa form of IL-1β (20k) appeared using the cotreatment with ATP and LPC/SPC (A and B). LPC/SPC inhibited ATP-induced mIL-1β release (A) and production of the p10 fragment (C) in a dose-dependent manner. If caspase-1 inhibitor (CaspI) was added to HBSS, the ATP-induced release of mIL-1β was inhibited showing the involvement of this protease in the maturation of IL-1β in MG6 cells (D). Treatment of the cells with 300 μM oATP during the LPS priming abolished the ATP-induced release of mIL-1β, suggesting the involvement of P2X7R, whereas the P2X7R function was not involved in LPC-induced release of pro- and mIL-1β (E). The asterisks in C indicate nonspecific bands corresponding to BSA contained in HBSS. The immunoblots are representative of at least three independent experiments. B, band intensities corresponding to mIL-1β (▪) and the 20-kDa form (□) in the supernatant from primary microglia were quantified in three independent experiments, and the data are indicated as a percentage of the amount of mIL-1β or the 20-kDa form obtained from the culture supernatant after the stimulation using 2 mM ATP in each set of experiments. The data are expressed as the mean value ± SEM. ∗, p < 0.05; ∗∗, p < 0.01; and ∗∗∗, p < 0.001.

Close modal

Similar to LPC, SPC alone induced the release of pro-IL-1β in a dose-dependent manner where a faint release of mIL-1β was detected at 80–100 μM and was abolished by the simultaneous treatment with 1 mM ATP in LPS-primed MG6 cells and primary microglia (Fig. 2,A). SPC also inhibits ATP-induced release of mIL-1β with a higher potency than LPC as observed using ELISA (Figs. 1 and 2,A). When primary microglia were treated with ATP at 1 and 2 mM, SPC promoted the release of the 20-kDa form of IL-1β at ∼40–60 μM (Fig. 2,A). However, the band corresponding to the 20-kDa form disappeared from the cells cotreated with 80–100 μM SPC. This indicates that SPC has the ability to inhibit both mature and the 20-kDa form of IL-1β in primary microglia. The immunoblot of the cell lysates indicated that the amounts of pro-IL-1β synthesized and stored within MG6 cells after priming with LPS were similar in each set of experiments (Fig. 2,A). mIL-1β was faintly detected in cell lysates using immunoblot due to the prompt release into the culture supernatant after processing (Fig. 2 A).

Using a densitometric analyses of the bands corresponding to the mature and the 20-kDa forms of IL-1β, mIL-1β release induced by 2 mM ATP was significantly inhibited by cotreatment with 60 μM LPC or 40 μM SPC in primary microglia (Fig. 2,B). Conversely, these concentrations of LPC and SPC promoted significant production of the 20-kDa form of IL-1β by stimulation with 2 mM ATP (Fig. 2 B).

Because caspase-1 plays a key role in the production of mIL-1β, we determined the effect of LPC and SPC on the activation of this enzyme in LPS-primed MG6 cells and primary microglia. The activated caspase-1 was assayed by determining the concentration of its processed 10-kDa subunit (p10) using immunoblot (29). The 45-kDa procaspase-1 was primarily detected in the cell lysates (Fig. 2,C). Both LPC and SPC promoted the release of procaspase-1 from microglial cells into the culture supernatant, but did not increase the amount of activated caspase-1 p10 in the culture supernatant as well as the cell lysates in MG6 cells (Fig. 2,C). Since LPC or SPC alone induced a release of mIL-1β weakly in MG6 cells, we speculate that the activation of caspase-1 induced by these lysophospholipids was weak and failed to be detected using immunoblot. In primary microglia, LPC was able to generate the p10 fragment of activated caspase-1 in the culture supernatant and cell lysates (Fig. 2,C). This indicates that primary microglia were more sensitive to LPC-induced activation with caspase-1 than MG6 cells. The treatment of microglial cells with 2 mM ATP increased the concentration of the p10 fragment in both the cell lysates and culture supernatant (Fig. 2,C). The increase in the amount of the p10 fragment induced by 2 mM ATP was inhibited using LPC or SPC in a dose-dependent manner (Fig. 2 C). This suggests that both LPC and SPC suppress ATP-induced mIL-1β release by interfering with caspase-1 activation.

The caspase-1 inhibitor Ac-YVAD-CMK blocked the release of mIL-1β induced by 2 mM ATP (Fig. 2,D), indicating the involvement of caspase-1 activation in ATP-induced mIL-1β release. Pretreatment with oATP inhibited the release of mIL-1β induced by 2 mM ATP, but did not affect the LPC-induced release of pro-IL-1β and mIL-1β (Fig. 2 E). This suggests that the activation of P2X7R is necessary for ATP-induced release of mIL-1β, but not for LPC-induced release of pro- and mIL-1β in MG6 cells.

The lysophospholipids LPC and SPC interact with the cell membrane due to their amphiphilic characters (30) and ultimately result in cell lysis. As expected, both LPC and SPC induced LDH leakage, a marker of cell damage, from LPS-primed MG6 cells at a concentration range of 60–100 μM (Fig. 3,A). This suggests the release of pro-IL-1β and procaspase-1 induced by LPC and SPC was due to their cell lytic actions. And possibly the lytic signals caused by LPC and SPC may result in caspase-1 activation followed by the maturation of IL-1β in MG6 cells (11). ATP induced a slight increase in extracellular LDH activity at 2 mM (Fig. 3,A). ATP suppressed the LDH leakage caused by LPC and SPC (Fig. 3,A). This suggests that ATP protects MG6 cells from lysophospholipid-induced cell damage. The suppressive effect on the leakage of LDH by ATP was more potent using LPC as compared with SPC stimulation (Fig. 3,A). ATP also suppressed the release of pro-IL-1β induced by LPC or SPC in MG6 cells (Fig. 3,B) and primary microglia (Fig. 3,C). ATP suppressed the LPC-induced pro-IL-1β release more efficiently than cells induced with SPC (Fig. 3, B and C). These data further support the correlation of LPC/SPC-induced release of pro-IL-1β with their lytic actions. Additionally, ATP significantly attenuated mIL-1β release that was induced by 60–100 μM LPC (Figs. 2,A and 3,B) and 100 μM SPC (Fig. 2,A) in MG6 cells. ATP also dose-dependently inhibited the release of mIL-1β induced by LPC and SPC in primary microglia, whereas the 20-kDa form of IL-1β appeared in LPC-treated primary cells according to the increased doses of ATP (Fig. 3 C). These findings suggest that ATP and the lysophospholipids mutually inhibited the production and release of mIL-1β in microglial cells.

FIGURE 3.

ATP inhibits LPC/SPC-induced LDH leakage and the release of pro- and mIL-1β in LPS-primed microglial cells. MG6 cells and primary microglia (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were incubated for 30 min in HBSS containing the test reagents at the concentrations indicated. The culture supernatant was collected and the cells were lysed with HBSS containing 1% Triton X-100. The activity of LDH in supernatant and lysates from MG6 cells was measured as described in Materials and Methods. LPC/SPC-induced LDH leakage in a dose-dependent manner was suppressed using the coapplication of ATP (A). The data are expressed as a percentage of the total cytosolic LDH obtained from nontreated MG6 cell lysates. Three independent experiments were performed and the data are expressed as the mean value ± SEM. ∗, p < 0.05; ∗∗, p < 0.01; and ∗∗∗, p < 0.001. B and C, Equal volumes of culture supernatant (sup) were loaded in each lane for SDS-PAGE. After transfer to a PVDF membrane, the blots were probed with anti-IL-1β Ab. LPC/SPC induced the release of pro-IL-1β (pro) and faint mIL-1β (m) that was inhibited using coapplication with ATP in MG6 cells (B) and primary microglia (C). Immunoblots are representative of at least three independent experiments.

FIGURE 3.

ATP inhibits LPC/SPC-induced LDH leakage and the release of pro- and mIL-1β in LPS-primed microglial cells. MG6 cells and primary microglia (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were incubated for 30 min in HBSS containing the test reagents at the concentrations indicated. The culture supernatant was collected and the cells were lysed with HBSS containing 1% Triton X-100. The activity of LDH in supernatant and lysates from MG6 cells was measured as described in Materials and Methods. LPC/SPC-induced LDH leakage in a dose-dependent manner was suppressed using the coapplication of ATP (A). The data are expressed as a percentage of the total cytosolic LDH obtained from nontreated MG6 cell lysates. Three independent experiments were performed and the data are expressed as the mean value ± SEM. ∗, p < 0.05; ∗∗, p < 0.01; and ∗∗∗, p < 0.001. B and C, Equal volumes of culture supernatant (sup) were loaded in each lane for SDS-PAGE. After transfer to a PVDF membrane, the blots were probed with anti-IL-1β Ab. LPC/SPC induced the release of pro-IL-1β (pro) and faint mIL-1β (m) that was inhibited using coapplication with ATP in MG6 cells (B) and primary microglia (C). Immunoblots are representative of at least three independent experiments.

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Previous studies demonstrate P2X7R-mediated release of mIL-1β from monocytes or macrophages is dependent on the K+ efflux from cells (12, 13). The K+ efflux results in the loss of cytosolic K+ and then triggers caspase-1 activation. In MG6 cells, the higher concentrations of extracellular K+ that directly inhibit K+ efflux from the cells completely blocked the ability of ATP to induce processing of IL-1β (data not shown). Nigericin, a K+/H+ ionophore, is known to induce processing of IL-1β in monocytes or macrophages using the efflux of K+ similar to the P2X7R-mediated mechanisms (31, 32). Therefore, we tested whether LPC and SPC inhibited nigericin-induced release of mIL-1β from LPS-primed MG6 cells. Nigericin promoted the release of mIL-1β from MG6 cells at a concentration of 1–10 μM (Fig. 4, A and B). We found that the nigericin-induced mIL-1β release was also blocked by the addition of higher concentrations of extracellular K+ (data not shown). LPC (Fig. 4,A) or SPC (Fig. 4,B) slightly enhanced the release of mIL-1β induced by 1 μM nigericin. Both lysophospholipids did not show significant inhibitory effects against the mIL-1β release induced by 5 and 10 μM nigericin (Fig. 4, A and B). These data suggest that the inhibitory mechanisms in mIL-1β release by LPC and SPC on P2X7R-mediated response may not be related to K+ efflux.

FIGURE 4.

K+ efflux, Ca2+ influx, and p44/42 MAPK activation are not associated with the inhibition of ATP-induced mIL-1β release using LPC and SPC in LPS-primed MG6 cells. MG6 cells (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 30 min in HBSS containing nigericin and LPC (A) or SPC (B) at the concentrations indicated. Equal volumes of culture supernatant (sup) were loaded in each lane for SDS-PAGE. After transfer to a PVDF membrane, the blots were probed with anti-IL-1β Ab. LPC/SPC did not inhibit the nigericin-induced release of mIL-1β (m) (A and B). The change in [K+]i was monitored using PBFI fluorescence as described in Materials and Methods. The fluorescence ratio of F340:F380 was used to estimate the change in [K+]i (C and D). The cells suspended in HBSS were treated with ATP and LPC (C) or SPC (D) at the concentrations indicated. LPC/SPC did not inhibit an ATP-induced decrease in [K+]i (C and D). MG6 cells cultured in 24-well plates were also incubated for 30 min in HBSS containing ATP in the presence or absence of extracellular Ca2+ (E, upper gels), A23187 (E, lower gels), and PD98059 (F) at the concentrations indicated. Equal volumes of culture supernatant were applied for SDS-PAGE and analyzed by immunoblot to detect IL-1β. Two millimolar ATP produced mIL-1β in the absence of extracellular Ca2+ (E, upper gels). Extracellular Ca2+ at >4 mM blocked the ATP-induced mIL-1β release due to the direct inhibition of P2X7R functions (E, upper gels). A23187 (A23) did not inhibit the ATP-induced mIL-1β release (E, lower gels). PD98059 (PD) showed no effect on the inhibition of ATP-induced mIL-1β release by LPC and SPC (F). DMSO (0.1%) was used as a control because PD98059 dissolves in DMSO (F). Immunoblots and traces of the fluorescence ratio of F340:F380 are representative of at least three independent experiments.

FIGURE 4.

K+ efflux, Ca2+ influx, and p44/42 MAPK activation are not associated with the inhibition of ATP-induced mIL-1β release using LPC and SPC in LPS-primed MG6 cells. MG6 cells (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 30 min in HBSS containing nigericin and LPC (A) or SPC (B) at the concentrations indicated. Equal volumes of culture supernatant (sup) were loaded in each lane for SDS-PAGE. After transfer to a PVDF membrane, the blots were probed with anti-IL-1β Ab. LPC/SPC did not inhibit the nigericin-induced release of mIL-1β (m) (A and B). The change in [K+]i was monitored using PBFI fluorescence as described in Materials and Methods. The fluorescence ratio of F340:F380 was used to estimate the change in [K+]i (C and D). The cells suspended in HBSS were treated with ATP and LPC (C) or SPC (D) at the concentrations indicated. LPC/SPC did not inhibit an ATP-induced decrease in [K+]i (C and D). MG6 cells cultured in 24-well plates were also incubated for 30 min in HBSS containing ATP in the presence or absence of extracellular Ca2+ (E, upper gels), A23187 (E, lower gels), and PD98059 (F) at the concentrations indicated. Equal volumes of culture supernatant were applied for SDS-PAGE and analyzed by immunoblot to detect IL-1β. Two millimolar ATP produced mIL-1β in the absence of extracellular Ca2+ (E, upper gels). Extracellular Ca2+ at >4 mM blocked the ATP-induced mIL-1β release due to the direct inhibition of P2X7R functions (E, upper gels). A23187 (A23) did not inhibit the ATP-induced mIL-1β release (E, lower gels). PD98059 (PD) showed no effect on the inhibition of ATP-induced mIL-1β release by LPC and SPC (F). DMSO (0.1%) was used as a control because PD98059 dissolves in DMSO (F). Immunoblots and traces of the fluorescence ratio of F340:F380 are representative of at least three independent experiments.

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To further assess the effect of LPC and SPC on [K+]i, the change in [K+]i was monitored using PBFI fluorescence (28). We observed that 5 mM ATP induced a sustained decrease in the F340:F380 ratio, indicating a continuous efflux of K+ from the cells (Fig. 4, C and D). Sixty micromolar LPC transiently decreased [K+]i (Fig. 4,C); and 60 μM SPC also induced a decrease in [K+]i, which gradually returned to the normal levels (Fig. 4,D). Coapplication of ATP and LPC/SPC additively promoted an efflux of [K+]i (Fig. 4, C and D). Collectively, the data support the inhibition of ATP-induced mIL-1β release by LPC and SPC was not due to the inhibition of K+ efflux from the cells.

We recently reported that LPC potentiates P2X7R-mediated intracellular signals in MG6 cells, i.e., Ca2+ influx and p44/42 MAPK activation (20). Therefore, the potentiation of Ca2+ influx and p44/42 MAPK activation may be associated with the inhibitory effects of LPC against ATP-induced mIL-1β release. Using 2 mM ATP induced mIL-1β release in the absence of extracellular Ca2+ (Fig. 4,E, upper gels). We previously showed that P2X7R-mediated Ca2+ influx was abolished in Ca2+-free HBSS (26), indicating that Ca2+ influx does not contribute to the ATP-induced maturation of IL-1β in MG6 cells. Extracellular Ca2+ at >4 mM blocked the ATP-induced mIL-1β release probably due to the direct inhibition of P2X7R functions (Fig. 4,E, upper gels) because P2X7R is allosterically antagonized by divalent cations such as Mg2+, Cu2+, and Ca2+ (33). Furthermore, a Ca2+ ionophore, A23187, that increases cytosolic Ca2+ levels failed to mimic the inhibitory effect of LPC and SPC on mIL-1β release induced by 2 mM ATP (Fig. 4 E, lower gels). Collectively, the data suggests that the inhibitory effect of LPC or SPC on ATP-induced mIL-1β release is not mediated by the potentiation of P2X7R-dependent Ca2+ influx.

Since LPC was demonstrated to potentiate the P2X7R-dependent p44/42 MAPK activation in MG6 cells (20), the up-regulated MAPK signals may correlate with the inhibitory effect of LPC on ATP-dependent mIL-1β release. However, the application of a MEK inhibitor, PD98059, at 50 μM, which was sufficient to inhibit the phosphorylation of p44/42 MAPK induced by 2 mM ATP in MG6 cells (data not shown), did not influence the inhibitory effects of LPC and SPC on ATP-induced mIL-1β release (Fig. 4 F). The data suggest that the activation of p44/42 MAPK was not associated with the inhibitory effects of both lysophospholipids.

We examined the subcellular distribution of pro-IL-1β in LPS-primed MG6 cells treated with or without 2 mM ATP. Pro-IL-1β was detected primarily in the cytosolic fraction (F1) in LPS-primed MG6 cells (Fig. 5,A). Trace amounts of pro-IL-1β were also detected in the membrane/organelle fraction (F2) (Fig. 5,A). In contrast, cathepsin D, a late endosome/early lysosome marker, was found in the membrane/organelle fraction (F2) (Fig. 5,A). There was no mIL-1β detected in any of the subcellular fractions in LPS-primed MG6 cells (Fig. 5,A). After stimulation with ATP at 2 mM for 30 min, the distribution of pro-IL-1β and cathepsin D did not change, and no significant increase in processed mIL-1β was detected in any of the subcellular fractions including F2 (Fig. 5,A). The data suggest that pro-IL-1β primarily exists as a cytosolic protein in LPS-primed MG6 cells and does not translocate to the lysosomal compartment for processing into the mature form as previously reported in macrophages (17). Using 2 mM ATP, LPS-primed MG6 cells developed numerous microvesicles that were eventually shed from the cell membranes (Fig. 5 B). It is likely that the shedding of microvesicles had entrapped the components of NALP3-inflammasome, known as a protein complex that stimulates caspase-1 activation and IL-1β maturation (34), and plays essential roles in the ATP-dependent release of mIL-1β in MG6 cells as shown in other microglial cell lines (19).

FIGURE 5.

Cytosolic distribution of pro-IL-1β and ATP-induced microvesicle formation and actin filament rearrangement in LPS-primed MG6 cells. MG6 cells (1 × 106 cells) were cultured in 35-mm dishes and further incubated with growth medium containing 1 μg/ml LPS for 4 h. Then, subcellular protein extracts were prepared from the cell monolayer as described in Materials and Methods. Each extract (16 μl) derived from four fractions, cytosol (F1), membrane/organelle (F2), nucleus (F3), and cytoskeleton (F4), was resolved using SDS-PAGE and immunoblot was performed using anti-IL-1β and anti-cathepsin D Abs (A). Pro-IL-1β was primarily detected in the cytosolic protein fraction (F1), whereas cathepsin D was found in the membrane/organelle fraction (F2) (A). The formation of microvesicles (▵) was observed in MG6 cells after treatment with 2 mM ATP (B). Additionally, MG6 cells were cultured in 8-well chamber slides (1 × 105 cells/well) and primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 10 min in HBSS containing 2 mM ATP. After fixation with formalin in PBS, the cells were immunostained with anti-actin Ab (C). The actin filament rearrangement (arrows) relates to the microvesicle shedding and was observed in ATP-treated cells but not in control cells (C). Immunoblots and microphotographs of immunostaining are representative of at least three independent experiments.

FIGURE 5.

Cytosolic distribution of pro-IL-1β and ATP-induced microvesicle formation and actin filament rearrangement in LPS-primed MG6 cells. MG6 cells (1 × 106 cells) were cultured in 35-mm dishes and further incubated with growth medium containing 1 μg/ml LPS for 4 h. Then, subcellular protein extracts were prepared from the cell monolayer as described in Materials and Methods. Each extract (16 μl) derived from four fractions, cytosol (F1), membrane/organelle (F2), nucleus (F3), and cytoskeleton (F4), was resolved using SDS-PAGE and immunoblot was performed using anti-IL-1β and anti-cathepsin D Abs (A). Pro-IL-1β was primarily detected in the cytosolic protein fraction (F1), whereas cathepsin D was found in the membrane/organelle fraction (F2) (A). The formation of microvesicles (▵) was observed in MG6 cells after treatment with 2 mM ATP (B). Additionally, MG6 cells were cultured in 8-well chamber slides (1 × 105 cells/well) and primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 10 min in HBSS containing 2 mM ATP. After fixation with formalin in PBS, the cells were immunostained with anti-actin Ab (C). The actin filament rearrangement (arrows) relates to the microvesicle shedding and was observed in ATP-treated cells but not in control cells (C). Immunoblots and microphotographs of immunostaining are representative of at least three independent experiments.

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Immunostaining shows the microvesicle shedding after treatment with 2 mM ATP was accompanied with the rearrangement of the actin filaments in the LPS-primed MG6 cells (Fig. 5,C). In the control cells (Fig. 6,A), rearrangement of actin filaments was not induced by treatment with 60 μM LPC (Fig. 6,B) or 40 μM SPC (Fig. 6,C) alone. MG6 cells treated with the lysophospholipids occasionally shrunk or exhibited a more rounded shape as compared with nontreated cells. The actin filament rearrangement related to the microvesicle shedding was observed in MG6 cells treated with 2 mM ATP (Fig. 6,D) and was blocked by the coadministration with LPC (Fig. 6,E) or SPC (Fig. 6 F).

FIGURE 6.

Effects of LPC and SPC on ATP-induced actin filament rearrangement in LPS-primed MG6 cells. MG6 cells were cultured in 8-well chamber slides (1 × 105 cells/well) and primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 10 min in HBSS containing the test reagents at the concentrations indicated. After fixation with formalin in PBS, the cells were immunostained with anti-actin Ab. In contrast to the control (A) and LPC/SPC-treated cells (B and C), the rearrangement of the actin filaments was clearly observed in the ATP-treated cells (D). The ATP-induced change in the actin filaments was suppressed by the cotreatment with LPC (E) or SPC (F). The microphotographs of immunostaining are representative of at least three independent experiments.

FIGURE 6.

Effects of LPC and SPC on ATP-induced actin filament rearrangement in LPS-primed MG6 cells. MG6 cells were cultured in 8-well chamber slides (1 × 105 cells/well) and primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 10 min in HBSS containing the test reagents at the concentrations indicated. After fixation with formalin in PBS, the cells were immunostained with anti-actin Ab. In contrast to the control (A) and LPC/SPC-treated cells (B and C), the rearrangement of the actin filaments was clearly observed in the ATP-treated cells (D). The ATP-induced change in the actin filaments was suppressed by the cotreatment with LPC (E) or SPC (F). The microphotographs of immunostaining are representative of at least three independent experiments.

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To assess the changes in microtubule organization, the cells were immunostained with anti-β-tubulin Ab. In the untreated MG6 cells, bundles of microtubules were loosely distributed in the cytoplasm (Fig. 7,A). LPC or SPC alone changed the cell shape to a more round appearance and the microtubules stained with anti-β-tubulin Ab were primarily located in the periphery of these round cells (Fig. 7, B and C). In cells treated with 2 mM ATP, the loose bundles of microtubules were rearranged into thick bundles and were radially extended from the centrosome toward the periphery of the cells (Fig. 7,D). Similar to the effect on actin filaments, LPC and SPC prevented the rearrangement of microtubules induced by ATP (Fig. 7, E and F). Additionally, the morphology of the cells cotreated with ATP and the lysophospholipids exhibited a more flattened attachment to the culture dishes as compared with the cells treated with LPC or SPC alone (Fig. 7, B, C, E, and F). If MG6 cells were pretreated with oATP, the radial rearrangement of microtubules induced by ATP was suppressed (Fig. 7 G), indicating that the microtubule rearrangement was mediated by P2X7R activation.

FIGURE 7.

The effects of LPC and SPC on ATP-induced microtubule reorganization in LPS-primed MG6 cells. MG6 cells were cultured in 8-well chamber slides (1 × 105 cells/well) and primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 10 min in HBSS containing the test reagents at the concentrations indicated. After fixation with formalin in PBS, the cells were immunostained with anti-β-tubulin Ab. In contrast to the control (A) and LPC/SPC-treated cells (B and C), the rearrangement of microtubules was clearly observed in the ATP-treated cells (D). The formation of radially extended bundles of microtubules induced by ATP was significantly suppressed using cotreatment with LPC (E) and SPC (F). Pretreatment with oATP, a P2X7R antagonist, inhibited the ATP-induced rearrangement of the microtubule bundles (G). Nigericin did not induce microtubule reorganization (H) and LPC showed no effect on nigericin-treated cells (I). The microphotographs of immunostaining are representative of at least three independent experiments.

FIGURE 7.

The effects of LPC and SPC on ATP-induced microtubule reorganization in LPS-primed MG6 cells. MG6 cells were cultured in 8-well chamber slides (1 × 105 cells/well) and primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 10 min in HBSS containing the test reagents at the concentrations indicated. After fixation with formalin in PBS, the cells were immunostained with anti-β-tubulin Ab. In contrast to the control (A) and LPC/SPC-treated cells (B and C), the rearrangement of microtubules was clearly observed in the ATP-treated cells (D). The formation of radially extended bundles of microtubules induced by ATP was significantly suppressed using cotreatment with LPC (E) and SPC (F). Pretreatment with oATP, a P2X7R antagonist, inhibited the ATP-induced rearrangement of the microtubule bundles (G). Nigericin did not induce microtubule reorganization (H) and LPC showed no effect on nigericin-treated cells (I). The microphotographs of immunostaining are representative of at least three independent experiments.

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Nigericin triggered the release of mIL-1β from the MG6 cells similar to ATP (Fig. 4, A and B). However, LPC and SPC suppressed the release of mIL-1β induced by ATP (Fig. 2, A and B) but not by nigericin (Fig. 4, A and B). With regard to the different effects of lysophospholipids among ATP- and nigericin-induced mIL-1β release, we observed that nigericin did not induce the radial extension of microtubule bundles observed in ATP-treated cells (Fig. 7,H). LPC also showed no effect on microtubule organization in nigericin-treated cells (Fig. 7 I). These data suggest that ATP-induced mIL-1β release is partially dependent on the state of microtubule organization, and the lysophospholipids are able to suppress the release of mIL-1β through the modulation of microtubule reorganization induced by ATP. We speculate that lysophospholipids do not inhibit the nigericin-induced mIL-1β release since the induction of mIL-1β release is independent of the state of microtubule organization.

Rho and Rho effectors are known to be involved in the formation of microtubules and the actin cytoskeleton (35). ATP is shown to induce the activation of RhoA using P2X7R in BAC1 macrophages (36). Therefore, we determined whether LPC/SPC modulates Rho-dependent pathways to suppress the cytoskeletal rearrangement induced by ATP in MG6 cells. In the presence of serum, RhoA was constitutively activated in MG6 cells regardless of priming with LPS (Fig. 8,A and data not shown). The basal activation level of RhoA probably reached the maximum in serum-containing medium because application of 2 mM ATP failed to induce additional activation of RhoA (Fig. 8,A). In contrast, the activation level of RhoA was low in the cells cultured in serum-free medium (Fig. 8,A). We found that in serum-free cultures 2 mM ATP increased the active form of RhoA (Fig. 8,A) due to the activation of P2X7R as described in a previous report (36). LPC or SPC alone suppressed the level of constitutively activated RhoA in both serum-containing and serum-free conditions (Fig. 8,A). In addition, LPC and SPC effectively decreased the levels of active RhoA even after stimulation of the cells with 2 mM ATP (Fig. 8,A). This suggests that these lysophospholipids likely down-regulate the ATP-induced cellular response by inhibiting RhoA activation. SPC was a more potent inhibitor of RhoA activation than LPC (Fig. 8,A) as observed with the inhibition of mIL-1β release (Figs. 1 and 2 A). This suggests that the RhoA-dependent pathway may play key roles in the process of ATP-induced cytoskeletal rearrangements and the release of mIL-1β in MG6 cells and LPC/SPC exhibits inhibitory effects by interfering with this pathway.

FIGURE 8.

LPC/SPC depresses RhoA activation in ATP-stimulated MG6 cells and ROCK inhibitors suppress ATP-induced mIL-1β release. MG6 cells (4 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 30 min in HBSS containing the test reagents at the concentrations indicated. In another experiment, the cells were cultured in serum-free medium and stimulated using HBSS containing the test reagents for 30 min. RhoA activation was measured using a pull-down assay with rhotekin beads as described in Materials and Methods. RhoA was constitutively activated in LPS-primed cells cultured in serum-containing medium that probably reached the plateau level because ATP failed to exert an additive effect (A). When MG6 cells were cultured in serum-free medium, the level of constitutively activated RhoA was reduced and ATP promoted the activation of RhoA (A). LPC and SPC depressed the constitutive activation of RhoA regardless of the presence of serum or LPS (A). After the stimulation with ATP, the level of active RhoA was still low in the cells cotreated with LPC or SPC (A). Two hundred micromolar GTPγS was used as a positive control for the detection of active RhoA (A). Band intensities corresponding to active RhoA were quantified in three independent experiments, and the data are indicated as a percentage of the maximum amount of active RhoA obtained from the cells stimulated by 2 mM ATP in each set of experiments (A). Additionally, MG6 cells (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h in the presence or absence of the ROCK inhibitors Y27632 and fasudil. After washing with PBS, the cells were further incubated for 30 min in HBSS containing ATP and the ROCK inhibitors at the concentrations indicated. The culture supernatant was collected and the cells were lysed with RIPA buffer. Equal volumes of supernatant (sup) or cell lysates were loaded in each lane for SDS-PAGE. After transfer to a PVDF membrane, the blots were probed with anti-IL-1β Ab (B). Y27632 and fasudil inhibited the ATP-induced release of mIL-1β (m) without affecting the cytosolic accumulation of pro-IL-1β (pro) in LPS-primed MG6 cells (B). The band intensities corresponding to mIL-1β in the supernatant were quantified in three independent experiments, and the data are shown as a percentage of the maximum amount of mIL-1β obtained from the culture supernatant after stimulation by 2 mM ATP in each set of experiments (B). The data shown are expressed as the mean value ± SEM. ∗, p < 0.05 and ∗∗, p < 0.01. Immunoblots are representative of three independent experiments.

FIGURE 8.

LPC/SPC depresses RhoA activation in ATP-stimulated MG6 cells and ROCK inhibitors suppress ATP-induced mIL-1β release. MG6 cells (4 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 30 min in HBSS containing the test reagents at the concentrations indicated. In another experiment, the cells were cultured in serum-free medium and stimulated using HBSS containing the test reagents for 30 min. RhoA activation was measured using a pull-down assay with rhotekin beads as described in Materials and Methods. RhoA was constitutively activated in LPS-primed cells cultured in serum-containing medium that probably reached the plateau level because ATP failed to exert an additive effect (A). When MG6 cells were cultured in serum-free medium, the level of constitutively activated RhoA was reduced and ATP promoted the activation of RhoA (A). LPC and SPC depressed the constitutive activation of RhoA regardless of the presence of serum or LPS (A). After the stimulation with ATP, the level of active RhoA was still low in the cells cotreated with LPC or SPC (A). Two hundred micromolar GTPγS was used as a positive control for the detection of active RhoA (A). Band intensities corresponding to active RhoA were quantified in three independent experiments, and the data are indicated as a percentage of the maximum amount of active RhoA obtained from the cells stimulated by 2 mM ATP in each set of experiments (A). Additionally, MG6 cells (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h in the presence or absence of the ROCK inhibitors Y27632 and fasudil. After washing with PBS, the cells were further incubated for 30 min in HBSS containing ATP and the ROCK inhibitors at the concentrations indicated. The culture supernatant was collected and the cells were lysed with RIPA buffer. Equal volumes of supernatant (sup) or cell lysates were loaded in each lane for SDS-PAGE. After transfer to a PVDF membrane, the blots were probed with anti-IL-1β Ab (B). Y27632 and fasudil inhibited the ATP-induced release of mIL-1β (m) without affecting the cytosolic accumulation of pro-IL-1β (pro) in LPS-primed MG6 cells (B). The band intensities corresponding to mIL-1β in the supernatant were quantified in three independent experiments, and the data are shown as a percentage of the maximum amount of mIL-1β obtained from the culture supernatant after stimulation by 2 mM ATP in each set of experiments (B). The data shown are expressed as the mean value ± SEM. ∗, p < 0.05 and ∗∗, p < 0.01. Immunoblots are representative of three independent experiments.

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Activated RhoA further induces activation of the Rho effector kinase (ROCK). We used two differing ROCK inhibitors, Y-27632 and fasudil, to determine whether the RhoA/ROCK pathway correlates with the ATP-induced release of mIL-1β in MG6 cells. Both inhibitors showed significant reduction of mIL-1β release induced by 2 mM ATP in a dose-dependent manner (Fig. 8 B). Collectively, the data suggest that the RhoA/ROCK pathway may play important roles in the ATP-induced release of mIL-1β and this pathway may be possibly affected by LPC and SPC.

To determine whether the modulation of microtubule rearrangement affects the ATP-induced mIL-1β release in MG6 cells, we used the microtubule inhibitor colchicine. Colchicine inhibits microtubule polymerization by binding to tubulin and is frequently used as an anti-inflammatory drug for the treatment of autoinflammatory diseases, including gout (37). Because pretreatment with this drug during LPS priming affects the cytosolic accumulation of pro-IL-1β in MG6 cells (data not shown), the cells were treated with colchicine combined with 2 mM ATP in HBSS for 30 min after LPS priming. Despite this short treatment period, colchicine dose-dependently suppressed the ATP-induced release of mIL-1β (Fig. 9,A). Although the radially extended microtubule bundles were stained with anti-β-tubulin Ab and was clearly observed in MG6 cells after stimulation with ATP, cotreatment with colchicine interfered with the formation of the radial extension of the microtubule bundles (Fig. 9 B). This suggests that modulation of microtubule rearrangement is associated with the ATP-induced mIL-1β release from MG6 cells. It also implies that LPC/SPC may affect the microtubule rearrangement process and inhibit ATP-induced release of mIL-1β in MG6 cells.

FIGURE 9.

Colchicine (col) inhibits ATP-induced mIL-1β release and microtubule reorganization in LPS-primed MG6 cells. MG6 cells (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 30 min in HBSS containing ATP and colchicine at the concentrations indicated. The culture supernatant was collected and the cells were lysed with RIPA buffer. Equal volumes of supernatant (sup) or cell lysates were loaded in each lane for SDS-PAGE. After transfer to a PVDF membrane, the blots were probed with anti-IL-1β Ab (A). The ATP-induced release of mIL-1β (m) was dose-dependently inhibited by treatment with colchicine (A). Band intensities corresponding to mIL-1β in supernatant were quantified in three independent experiments, and the data are indicated as a percentage of the maximum amount of mIL-1β obtained from the culture supernatant after the stimulation by 2 mM ATP in each set of experiments (A). The data are expressed as the mean value ± SEM. ∗, p < 0.05. Additionally, MG6 cells were cultured in 8-well chamber slides (1 × 105 cells/well) and primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 10 min in HBSS containing ATP and colchicine at the concentrations indicated. After fixation with formalin in PBS, the cells were immunostained with anti-β-tubulin Ab. In contrast to the control and the colchicine-treated cells, the radially extended thick bundles of microtubule were observed in ATP-treated cells (B). Colchicine inhibited the ATP-induced microtubule reorganization (B). The microphotographs of immunostaining are representative of at least three experiments.

FIGURE 9.

Colchicine (col) inhibits ATP-induced mIL-1β release and microtubule reorganization in LPS-primed MG6 cells. MG6 cells (3 × 105 cells/well in 24-well plates) were primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 30 min in HBSS containing ATP and colchicine at the concentrations indicated. The culture supernatant was collected and the cells were lysed with RIPA buffer. Equal volumes of supernatant (sup) or cell lysates were loaded in each lane for SDS-PAGE. After transfer to a PVDF membrane, the blots were probed with anti-IL-1β Ab (A). The ATP-induced release of mIL-1β (m) was dose-dependently inhibited by treatment with colchicine (A). Band intensities corresponding to mIL-1β in supernatant were quantified in three independent experiments, and the data are indicated as a percentage of the maximum amount of mIL-1β obtained from the culture supernatant after the stimulation by 2 mM ATP in each set of experiments (A). The data are expressed as the mean value ± SEM. ∗, p < 0.05. Additionally, MG6 cells were cultured in 8-well chamber slides (1 × 105 cells/well) and primed with 1 μg/ml LPS for 4 h. After washing with PBS, the cells were further incubated for 10 min in HBSS containing ATP and colchicine at the concentrations indicated. After fixation with formalin in PBS, the cells were immunostained with anti-β-tubulin Ab. In contrast to the control and the colchicine-treated cells, the radially extended thick bundles of microtubule were observed in ATP-treated cells (B). Colchicine inhibited the ATP-induced microtubule reorganization (B). The microphotographs of immunostaining are representative of at least three experiments.

Close modal

In this study, we demonstrated for the first time that LPC and SPC have the ability to suppress the release and processing of IL-1β in microglial cells upon stimulation with ATP. These lysophospholipids blocked caspase-1 activation, which is required for IL-1β maturation. ATP induced dynamic rearrangement of the actin and tubulin cytoskeleton using P2X7R activation in MG6 cells and LPC and SPC blocked these cytoskeletal changes. LPC/SPC reduced the active form of RhoA and ROCK inhibitors prevent the ATP-induced release of mIL-1β. Therefore, RhoA-dependent cytoskeletal reorganization may have essential roles in the inhibitory actions of LPC/SPC on ATP-induced maturation of IL-1β. Additionally, the microtubule inhibitor colchicine inhibited the ATP-induced release of mIL-1β, suggesting the involvement of microtubule reorganization in the IL-1β maturation in microglia. Collectively, we suggest a model for the roles of LPC/SPC in the inhibition of P2X7R-dependent mIL-1β release in microglial cells (Fig. 10, pathway 1).

FIGURE 10.

Schematic model for the mutual regulatory mechanism of IL-1β maturation by lysophospholipids and ATP in microglial cells. In pathway 1, LPC/SPC likely modulates RhoA-dependent cytoskeletal rearrangement (orange square) and suppresses P2X7R-dependent caspase-1 activation (□) that is required for the maturation of IL-1β (blue square). In pathway 2, the cytoskeletal rearrangements induced by P2X7R activation (orange square) suppress the LPC/SPC-dependent LDH leakage and release of pro-IL-1β (purple square) that may be possibly mediated by a local mechanical stress through the direct insertion of these lysophospholipids into the lipid bilayer of the cell membrane (yellow square). In addition, the P2X7R-mediated suppression of LPC/SPC-dependent lytic actions may contribute to the inhibition of LPC/SPC-dependent caspase-1 activation (□) and IL-1β maturation (blue square).

FIGURE 10.

Schematic model for the mutual regulatory mechanism of IL-1β maturation by lysophospholipids and ATP in microglial cells. In pathway 1, LPC/SPC likely modulates RhoA-dependent cytoskeletal rearrangement (orange square) and suppresses P2X7R-dependent caspase-1 activation (□) that is required for the maturation of IL-1β (blue square). In pathway 2, the cytoskeletal rearrangements induced by P2X7R activation (orange square) suppress the LPC/SPC-dependent LDH leakage and release of pro-IL-1β (purple square) that may be possibly mediated by a local mechanical stress through the direct insertion of these lysophospholipids into the lipid bilayer of the cell membrane (yellow square). In addition, the P2X7R-mediated suppression of LPC/SPC-dependent lytic actions may contribute to the inhibition of LPC/SPC-dependent caspase-1 activation (□) and IL-1β maturation (blue square).

Close modal

LPC is known as a proinflammatory lipid that is generated by hydrolysis of phosphatidylcholine, a major constituent of the plasma membrane, using the function of phospholipase A2 (PLA2). LPC was recently reported to be a potent endogenous inducer of mIL-1β release in microglial cells (21). In agreement with this report, we demonstrated that LPC alone triggered the release of mIL-1β in mouse microglial cells. In primary microglia, the lower concentrations of LPC (20–40 μM) were able to generate a significant amount of mIL-1β, suggesting that this event physiologically occurs during inflammation as described in a previous report (21). In contrast, we demonstrated the anti-inflammatory function of LPC by showing the inhibition of P2X7R-dependent mIL-1β release. Because the inhibitory effect of LPC was observed at concentrations of ∼20–40 μM and these concentrations of LPC did not significantly affect the LDH leakage from MG6 cells, we consider the inhibitory action of LPC also has biological significance in vivo. During inflammation in the brain, ATP is released from damaged cells and LPC is produced by PLA2 and secreted extracellularly. Our data therefore suggest that LPC primarily acts in microglia as a proinflammatory factor; but later it plays anti-inflammatory roles by inhibiting P2X7R-dependent maturation of IL-1β and then protects the brain tissue from damage caused by prolonged inflammation.

Little is known about the formation and degradation of SPC and its cellular targets. Evidence suggests its possible role as a lipid mediator in a variety of cell types (38). In the CNS, SPC likely has some physiological roles because it induces a neurite outgrowth in neuroblastoma cell lines (39). Additionally, the massive brain dysfunction observed in Niemann-Pick disease type A is reported to be associated with the accumulation of SPC (40). This suggests that SPC may play important roles under both physiological and pathological conditions in the brain. Until now, there is no information showing the direct effect of SPC on microglial cells. However, among the lysophospholipids we tested, SPC showed the strongest inhibition of the P2X7R-dependent mIL-1β release in microglial cells. Thus, we speculate that SPC acts in the microglia as a modulator of the inflammatory response in the brain.

In the pathological brain, the production of LPC and SPC as well as the release of ATP from damaged cells to the extracellular space increases (40, 41). The function and expression of P2X7R in microglial cells is also reported to be up-regulated in several neurodegenerative disorders including Alzheimer’s disease and prion diseases (42, 43). Therefore, increased accumulation of these factors possibly impairs the physiological regulation of P2X7R-dependent mIL-1β release in microglial cells that may ultimately contribute to the onset of the neuron degeneration found in such brain diseases.

Recent studies suggest that the action of LPC and SPC may be mediated by G protein-coupled receptors, i.e., G2A and GPR4 (44). In microglial cells, we previously reported that modulation of P2X7R-mediated intracellular signals by LPC is independent of the G protein-coupled receptor-mediated pathways (20). This is also supported by a recent study showing that the potentiation of P2X7R-mediated functions by lysophospholipids, including LPC and SPC, is predominantly due to lipid-induced changes in membrane properties in the THP-1 cells because there was little structural specificity of the lipids to induce these effects (22). In this study, our data also show that other lysolipid molecules such as platelet-activating factor and edelfosine (ET-18-OCH3) exert the same inhibitory effect against ATP-induced mIL-1β release at similar concentrations (20–100 μM; data not shown). The common potency of different amphiphilic lysolipids implies the direct association of LPC and SPC with the plasma membrane, rather than the mediation by specific receptor systems, is responsible for their lytic actions as well as the inhibition of ATP-induced mIL-1β release in microglial cells.

Bianco et al. (19) first reported that vesicle shedding dominantly contributes to the ATP-induced release of mIL-1β in microglial cells (19). They also suggest a lesser contribution by the exocytotic pathway of lysosomal-related vesicles for mIL-1β release in microglial cells because a PLA2 blocker, AAC0CF3, failed to block the mIL-1β release from microglial cells. We observed an abundant formation of microvesicles shedding from the cell membrane in MG6 cells after ATP stimulation. Pro-IL-1β accumulated in the cytosolic fraction that is a different compartment than the cathepsin D-containing lysosomal fraction. These data suggest that microvesicle shedding rather than the lysosome exocytosis primarily contributes to the ATP-induced release of mIL-1β from MG6 cells as reported previously (19). The actin cytoskeleton is reported to be important for the formation of the membrane bleb and, subsequently, the vesicle shedding induced by P2X7R activation (45). In MG6 cells, we found actin rearrangement related to the formation of microvesicles after the treatment with ATP. Furthermore, we demonstrated the dynamic reorganization of microtubules induced by the P2X7R activation in MG6 cells. To our knowledge, this is the first report showing ATP-induced microtubule reorganization and its contribution to P2X7R-dependent mIL-1β release in microglial cells.

In addition to the 17-kDa mature form of IL-1β, we observed that the release of the 20-kDa fragment of IL-1β was induced by ATP in the presence of LPC or SPC in primary microglia. Although previous studies provided brief comments about a similar fragment of IL-1β that has a slightly higher molecular mass than mIL-1β (31, 46), its biological significance or detailed secretion mechanisms remain unknown. The production of the 20-kDa fragment may be under some physiological regulation in microglial cells because ATP combined with higher doses of LPC facilitated the generation of this fragment, and SPC stimulated the release of this fragment at 40 μM but it was inhibited at 60–100 μM. Thus, the in vitro microglial system may provide a useful model to further analyze the processing and release of this novel IL-1β fragment.

The effect of ROCK inhibitors on ATP-induced mIL-1β release in macrophages and microglia is apparently conflicting. The same ROCK inhibitors we used in this study failed to inhibit the ATP-induced release of mIL-1β in BAC1 macrophages (36). The discrepancy may be due to the difference in the levels of active RhoA in these cells. In BAC1 macrophages, active RhoA is not detected when the cells are cultured in serum-containing medium. In contrast, MG6 cells contained significantly higher amounts of active RhoA in the presence or absence of serum. Since the basal level of activated RhoA in BAC1 cells is low compared with MG6 cells, we speculate that different intrinsic regulation of RhoA activation reflects the different contribution of RhoA-dependent cytoskeletal changes in P2X7R-mediated mIL-1β release between BAC1 and MG6 cells.

Martinon et al. (47) demonstrated that colchicine blocks the release of mIL-1β induced by monosodium urate and calcium pyrophosphate dihydrate crystals but not by ATP in THP-1 cells (47). However, we observed that colchicine inhibited the ATP-induced mIL-1β release in MG6 cells. The ATP-stimulated pathway leading to mIL-1β release in mouse microglial MG6 cells may be different than in the human monocytic THP-1 cells. One possibility is the differential contributory ratio of vesicle shedding:lysosome exocytosis to ATP-induced mIL-1β release among these two cell lines. A recent finding shows that LPC and SPC enhance the ATP-induced IL-1β release in THP-1 cells (22) and may support our speculation that MG6 and THP-1 cells have different intrinsic pathways for P2X7R-dependent release of mIL-1β.

As shown in Fig. 10 (pathway 2), ATP conversely inhibited the LPC/SPC-induced LDH leakage and the release of pro-IL-1β in MG6 cells. LPC, as well as SPC, is an amphiphilic molecule that produces local mechanical stress or membrane damage by inserting itself into the outer monolayer of the plasma membrane (30). Because LDH leakage is well known as a marker of necrotic cell death and cell lysis, it may be that ATP protected the cell from damage induced by these lysophospholipids. We showed that the cells treated with higher concentrations of LPC or SPC (>60 μM) alone occasionally exhibited shrunken morphology, probably due to the loss of membrane integrity caused by the lytic actions. In contrast, the cells cotreated with ATP and LPC/SPC were healthy and kept their extended morphology attached onto the culture dishes. Such a difference in cell morphology leads us to speculate that P2X7R-dependent cytoskeletal reorganization modulates the LPC/SPC-dependent cell shrinkage. Similar to the mechanism for other injurious stimuli such as with crystalline silica (48), the lytic properties of LPC and SPC are partially responsible for the maturation of IL-1β in microglial cells because LDH leakage precedes the production of mIL-1β, especially in MG6 cells. Therefore, the inhibitory actions of ATP on the LPC/SPC-induced IL-1β maturation may be due to the suppression of their lytic properties using P2X7R-dependent rearrangement of the cytoskeleton in microglial cells. Further studies will be necessary to unravel the basis whereby ATP inhibits LPC/SPC-induced cell lysis in microglia.

In conclusion, this study proposes that ATP and lysophospholipids (LPC/SPC) have two opposite effects, proinflammatory and anti-inflammatory roles, in the immunological response in microglial cells. The mutual suppression of IL-1β maturation by lysophospholipids and ATP may have physiological significance in regulating the microglial function in the brain. With regard to anti-inflammatory effects, recent reports demonstrate the therapeutic effects of LPC/SPC on experimental sepsis through the suppression of inflammatory responses in vivo (23, 24, 25). In these studies, the administration of LPC/SPC significantly reduces the levels of IL-1β caused by endotoxemia. Our finding that LPC/SPC inhibits P2X7R-mediated IL-1β release in LPS-primed cells may explain the molecular mechanisms by which these lysophospholipids inhibit IL-1β production during experimental sepsis in vivo. Our MG6 microglial cell line will provide useful experimental systems to determine the molecular and biochemical mechanisms for the modulation of IL-1β maturation mediated by P2X7R.

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.

1

This work was supported by a research grant and Grant-in-Aid from the BSE Control Project of the Ministry of Agriculture, Forestry, and Fisheries in Japan, and a Grant-in-Aid for Young Scientists (Category B) from the Ministry of Education, Science, Sports and Technology of Japan. This work was also supported by a research grant from The Naito Foundation.

3

Abbreviations used in this paper: mIL-1β, mature IL-1β; LPC, lysophosphatidylcholine; SPC, sphingosylphosphorylcholine; PLA2, phospholipase A2; oATP, oxidized ATP; PVDF, polyvinylidene difluoride.

1
Streit, W. J..
2002
. Microglia as neuroprotective, immunocompetent cells of the CNS.
Glia
40
:
133
-139.
2
Nimmerjahn, A., F. Kirchhoff, F. Helmchen.
2005
. Resting microglial cells are highly dynamic surveillants of brain parenchyma in vivo.
Science
308
:
1314
-1318.
3
Hanisch, U. K..
2002
. Microglia as a source and target of cytokines.
Glia
40
:
140
-155.
4
Blum-Degen, D., T. Muller, W. Kuhn, M. Gerlach, H. Przuntek, P. Riederer.
1995
. Interleukin-1β and interleukin-6 are elevated in the cerebrospinal fluid of Alzheimer’s and de novo Parkinson’s disease patients.
Neurosci. Lett.
202
:
17
-20.
5
Lucas, S. M., N. J. Rothwell, R. M. Gibson.
2006
. The role of inflammation in CNS injury and disease.
Br. J. Pharmacol.
147
: (Suppl. 1):
S232
-S240.
6
Dinarello, C. A..
2002
. The IL-1 family and inflammatory diseases.
Clin. Exp. Rheumatol.
20
:
S1
-S13.
7
Dinarello, C. A..
1996
. Biologic basis for interleukin-1 in disease.
Blood
87
:
2095
-2147.
8
Yamin, T. T., J. M. Ayala, D. K. Miller.
1996
. Activation of the native 45-kDa precursor form of interleukin-1-converting enzyme.
J. Biol. Chem.
271
:
13273
-13282.
9
Sanz, J. M., F. Di Virgilio.
2000
. Kinetics and mechanism of ATP-dependent IL-1β release from microglial cells.
J. Immunol.
164
:
4893
-4898.
10
Andrei, C., C. Dazzi, L. Lotti, M. R. Torrisi, G. Chimini, A. Rubartelli.
1999
. The secretory route of the leaderless protein interleukin 1β involves exocytosis of endolysosome-related vesicles.
Mol. Biol. Cell
10
:
1463
-1475.
11
Wewers, M. D..
2004
. IL-1β: an endosomal exit.
Proc. Natl. Acad. Sci. USA
101
:
10241
-10242.
12
Andrei, C., P. Margiocco, A. Poggi, L. V. Lotti, M. R. Torrisi, A. Rubartelli.
2004
. Phospholipases C and A2 control lysosome-mediated IL-1β secretion: Implications for inflammatory processes.
Proc. Natl. Acad. Sci. USA
101
:
9745
-9750.
13
Kahlenberg, J. M., G. R. Dubyak.
2004
. Mechanisms of caspase-1 activation by P2X7 receptor-mediated K+ release.
Am. J. Physiol.
286
:
C1100
-C1108.
14
Gudipaty, L., J. Munetz, P. A. Verhoef, G. R. Dubyak.
2003
. Essential role for Ca2+ in regulation of IL-1β secretion by P2X7 nucleotide receptor in monocytes, macrophages, and HEK-293 cells.
Am. J. Physiol.
285
:
C286
-C299.
15
Ferrari, D., C. Pizzirani, E. Adinolfi, R. M. Lemoli, A. Curti, M. Idzko, E. Panther, F. Di Virgilio.
2006
. The P2X7 receptor: a key player in IL-1 processing and release.
J. Immunol.
176
:
3877
-3883.
16
MacKenzie, A., H. L. Wilson, E. Kiss-Toth, S. K. Dower, R. A. North, A. Surprenant.
2001
. Rapid secretion of interleukin-1β by microvesicle shedding.
Immunity
15
:
825
-835.
17
Brough, D., N. J. Rothwell.
2007
. Caspase-1-dependent processing of pro-interleukin-1β is cytosolic and precedes cell death.
J. Cell Sci.
120
:
772
-781.
18
Qu, Y., L. Franchi, G. Nunez, G. R. Dubyak.
2007
. Nonclassical IL-1β secretion stimulated by P2X7 receptors is dependent on inflammasome activation and correlated with exosome release in murine macrophages.
J. Immunol.
179
:
1913
-1925.
19
Bianco, F., E. Pravettoni, A. Colombo, U. Schenk, T. Moller, M. Matteoli, C. Verderio.
2005
. Astrocyte-derived ATP induces vesicle shedding and IL-1β release from microglia.
J. Immunol.
174
:
7268
-7277.
20
Takenouchi, T., M. Sato, H. Kitani.
2007
. Lysophosphatidylcholine potentiates Ca2+ influx, pore formation and p44/42 MAP kinase phosphorylation mediated by P2X7 receptor activation in mouse microglial cells.
J. Neurochem.
102
:
1518
-1532.
21
Stock, C., T. Schilling, A. Schwab, C. Eder.
2006
. Lysophosphatidylcholine stimulates IL-1β release from microglia via a P2X7 receptor-independent mechanism.
J. Immunol.
177
:
8560
-8568.
22
Michel, A. D., E. Fonfria.
2007
. Agonist potency at P2X7 receptors is modulated by structurally diverse lipids.
Br. J. Pharmacol.
152
:
523
-537.
23
Yan, J. J., J. S. Jung, J. E. Lee, J. Lee, S. O. Huh, H. S. Kim, K. C. Jung, J. Y. Cho, J. S. Nam, H. W. Suh, et al
2004
. Therapeutic effects of lysophosphatidylcholine in experimental sepsis.
Nat. Med.
10
:
161
-167.
24
Chen, G., J. Li, X. Qiang, C. J. Czura, M. Ochani, K. Ochani, L. Ulloa, H. Yang, K. J. Tracey, P. Wang, et al
2005
. Suppression of HMGB1 release by stearoyl lysophosphatidylcholine: an additional mechanism for its therapeutic effects in experimental sepsis.
J. Lipid. Res.
46
:
623
-627.
25
Murch, O., M. Abdelrahman, M. Collino, M. Gallicchio, E. Benetti, E. Mazzon, R. Fantozzi, S. Cuzzocrea, C. Thiemermann.
2008
. Sphingosylphosphorylcholine reduces the organ injury/dysfunction and inflammation caused by endotoxemia in the rat.
Crit. Care Med.
36
:
550
-559.
26
Takenouchi, T., K. Ogihara, M. Sato, H. Kitani.
2005
. Inhibitory effects of U73122 and U73343 on Ca2+ influx and pore formation induced by the activation of P2X7 nucleotide receptors in mouse microglial cell line.
Biochim. Biophys. Acta
1726
:
177
-186.
27
Suzumura, A., S. G. Mezitis, N. K. Gonatas, D. H. Silberberg.
1987
. MHC antigen expression on bulk isolated macrophage-microglia from newborn mouse brain: induction of Ia antigen expression by γ-interferon.
J. Neuroimmunol.
15
:
263
-278.
28
Galvan, E., M. Sitges.
2004
. Characterization of the participation of sodium channels on the rise in Na+ induced by 4-aminopyridine (4-AP) in synaptosomes.
Neurochem. Res.
29
:
347
-355.
29
Verhoef, P. A., S. B. Kertesy, K. Lundberg, J. M. Kahlenberg, G. R. Dubyak.
2005
. Inhibitory effects of chloride on the activation of caspase-1, IL-1β secretion, and cytolysis by the P2X7 receptor.
J. Immunol.
175
:
7623
-7634.
30
Lundbaek, J. A., O. S. Andersen.
1994
. Lysophospholipids modulate channel function by altering the mechanical properties of lipid bilayers.
J. Gen. Physiol.
104
:
645
-673.
31
Perregaux, D., J. Barberia, A. J. Lanzetti, K. F. Geoghegan, T. J. Carty, C. A. Gabel.
1992
. IL-1β maturation: evidence that mature cytokine formation can be induced specifically by nigericin.
J. Immunol.
149
:
1294
-1303.
32
Perregaux, D., C. A. Gabel.
1994
. Interleukin-1β maturation and release in response to ATP and nigericin: evidence that potassium depletion mediated by these agents is a necessary and common feature of their activity.
J. Biol. Chem.
269
:
15195
-15203.
33
Virginio, C., D. Church, R. A. North, A. Surprenant.
1997
. Effects of divalent cations, protons, and calmidazolium at the rat P2X7 receptor.
Neuropharmacology
36
:
1285
-1294.
34
Ogura, Y., F. S. Sutterwala, R. A. Flavell.
2006
. The inflammasome: first line of the immune response to cell stress.
Cell
126
:
659
-662.
35
Ishizaki, T., Y. Morishima, M. Okamoto, T. Furuyashiki, T. Kato, S. Narumiya.
2001
. Coordination of microtubules and the actin cytoskeleton by the Rho effector mDia1.
Nat. Cell. Biol.
3
:
8
-14.
36
Verhoef, P. A., M. Estacion, W. Schilling, G. R. Dubyak.
2003
. P2X7 receptor-dependent blebbing and the activation of Rho-effector kinases, caspases, and IL-1β release.
J. Immunol.
170
:
5728
-5738.
37
Molad, Y..
2002
. Update on colchicine and its mechanism of action.
Curr. Rheumatol. Rep.
4
:
252
-256.
38
Meyer zu Heringdorf, D., H. M. Himmel, K. H. Jakobs.
2002
. Sphingosylphosphorylcholine-biological functions and mechanisms of action.
Biochim. Biophys. Acta
1582
:
178
-189.
39
Sugiyama, E., K. Uemura, A. Hara, T. Taketomi.
1990
. Effects of various lysosphingolipids on cell growth, morphology and lipid composition in three neuroblastoma cell lines.
Biochem. Biophys. Res. Commun.
169
:
673
-679.
40
Rodriguez-Lafrasse, C., M. T. Vanier.
1999
. Sphingosylphosphorylcholine in Niemann-Pick disease brain: accumulation in type A but not in type B.
Neurochem. Res.
24
:
199
-205.
41
Farooqui, A. A., M. L. Litsky, T. Farooqui, L. A. Horrocks.
1999
. Inhibitors of intracellular phospholipase A2 activity: their neurochemical effects and therapeutical importance for neurological disorders.
Brain Res. Bull.
49
:
139
-153.
42
Takenouchi, T., Y. Iwamaru, M. Imamura, N. Kato, S. Sugama, M. Fujita, M. Hashimoto, M. Sato, H. Okada, T. Yokoyama, et al
2007
. Prion infection correlates with hypersensitivity of P2X7 nucleotide receptor in a mouse microglial cell line.
FEBS Lett.
581
:
3019
-3026.
43
McLarnon, J. G., J. K. Ryu, D. G. Walker, H. B. Choi.
2006
. Upregulated expression of purinergic P2X7 receptor in Alzheimer disease and amyloid-β peptide-treated microglia and in peptide-injected rat hippocampus.
J. Neuropathol. Exp. Neurol.
65
:
1090
-1097.
44
Xu, Y..
2002
. Sphingosylphosphorylcholine and lysophosphatidylcholine: G protein-coupled receptors and receptor-mediated signal transduction.
Biochim. Biophys. Acta
1582
:
81
-88.
45
Pfeiffer, Z. A., M. Aga, U. Prabhu, J. J. Watters, D. J. Hall, P. J. Bertics.
2004
. The nucleotide receptor P2X7 mediates actin reorganization and membrane blebbing in RAW 264.7 macrophages via p38 MAP kinase and Rho.
J. Leukocyte Biol.
75
:
1173
-1182.
46
Pelegrin, P., A. Surprenant.
2007
. Pannexin-1 couples to maitotoxin- and nigericin-induced interleukin-1β release through a dye uptake-independent pathway.
J. Biol. Chem.
282
:
2386
-2394.
47
Martinon, F., V. Petrilli, A. Mayor, A. Tardivel, J. Tschopp.
2006
. Gout-associated uric acid crystals activate the NALP3 inflammasome.
Nature
440
:
237
-241.
48
Schmidt, J. A., C. N. Oliver, J. L. Lepe-Zuniga, I. Green, I. Gery.
1984
. Silica-stimulated monocytes release fibroblast proliferation factors identical to interleukin 1: a potential role for interleukin 1 in the pathogenesis of silicosis.
J. Clin. Invest.
73
:
1462
-1472.