TLRs detect conserved molecular patterns that are unique to microbes, enabling tailored responses to invading pathogens and modulating a multitude of immunopathological conditions. We investigated the ability of a naturally occurring stearoyl-arachidonoyl form of phosphatidylserine (SAPS) to inhibit the proinflammatory effects of TLR agonists in models of inflammation investigating the interaction of leukocytes with epithelial and endothelial cells. The responses to LPS of both epithelial and endothelial cells were highly amplified in the presence of PBMCs. Coincubation with SAPS markedly inhibited activation of cocultures by LPS, principally through inhibition of the TLR4 signaling pathway in PBMCs; however, this was not through downmodulation of TLR4 or coreceptor expression, nor was IL-1β-induced cytokine release affected. SAPS also impaired Pam3CSK4 (TLR2/1), Gardiquimod (TLR7/8), and Streptococcus pneumoniae-induced cytokine release, but had only modest effects on poly(I:C) (TLR3)-induced responses. Fluorescence resonance energy transfer analysis of molecular associations revealed that SAPS disrupted the association of both TLR4 and TLR2 with their respective membrane partners that are required for signaling. Thus, our data reinforce the existence and importance of cooperative networks of TLRs, tissue cells, and leukocytes in mediating innate immunity, and identify a novel disrupter of membrane microdomains, revealing the dependence of TLR signaling on localization within these domains.

It is now well established that TLRs detect conserved molecular patterns unique to microbes, facilitating host defense against pathogens and enabling the construction of tailored responses that are essential for their clearance (1, 2). TLRs are key in the detection of many structurally unrelated proinflammatory stimuli, including a diverse range of microbial products and also endogenous ligands, generated from damaged or dying host cells (reviewed in Refs. 3 , 4).

TLR4 is the most widely studied member of the family and recognizes an extensive range of agonists, but primarily LPS (5, 6). LPS-induced cellular activation is thought to occur when receptors are activated within membrane microdomains such as lipid rafts, which may help to confer ligand specificity through the association of diverse proteins within the raft (7, 8). CD14 is a key protein within rafts and is involved in transfer of LPS to the TLR4/MD-2 complex (9, 10, 11). CD14 also binds to, and mediates the transport of, phospholipids such as phosphatidylserine (PS)3 (12, 13, 14, 15). While phosphatidylinositol has been reported to be an LPS antagonist (13, 16), PS has been shown to have only modest effects on LPS signaling, potentially as a result of interference with the LPS/CD14 interaction (16), although the exact mechanism remains unclear. PS may also have other effects on TLR signaling. It is ubiquitously present in the mammalian cells, where it is normally located on the cytosolic surface of the plasma membrane (17). During apoptosis, PS is externalized to the outer membrane, targeting the cell for recognition and clearance by macrophages (18, 19). This process of apoptotic clearance leads to the production of antiinflammatory cytokines and the active suppression of inflammatory mediator production (20, 21). While PS is important in recognition of apoptotic cells, it is less clear whether PS directly signals to drive the phenotype of engulfing macrophages to a more antiinflammatory state.

We were therefore interested to explore the consequences of PS exposure on subsequent TLR responses in a range of cell types and in vitro coculture models. We have previously demonstrated that responses to TLR agonists are often most efficaciously induced when leukocytes and tissue cells are allowed to interact, and proposed that tissue cell signaling is an important amplification mechanism for signals derived from leukocytes interacting with microbial agonists (22, 23, 24, 25). In previous studies we have shown that the neutralization of IL-1 can inhibit these networks (23). In this study we observed that a naturally occurring species of PS, 1-stearoyl-2-arachidonoyl-sn-glycero-3-[phospho-l-serine] (SAPS), was an effective inhibitor of TLR4, but not IL-1β, signaling. Its nontoxic mode of action was not likely to be due to competition for LPS or activation of antiinflammatory pathways, but was consistent with its ability to disrupt membrane microdomains, and revealed a substantial dependence of many TLRs on such domains for effective signaling.

Cell culture reagents were purchased from Invitrogen, and general laboratory reagents were purchased from Sigma-Aldrich. FCS (endotoxin levels <0.5 EU/ml) was purchased from BioWhittaker. Purified LPS from Escherichia coli serotype R515 was from Axxora. The synthetic lipopeptide Pam3CysSerLys4 (Pam3CSK4) was from EMC Microcollections (Tübingen, Germany). Poly(I:C) and Gardiquimod were from InvivoGen. Recombinant human IL-1β and TNF-α were from PeproTech. Inert (heat-killed) type 2 Streptococcus pneumoniae (strain D39) was a kind gift from Dr. D. H. Dockrell (26). Highly purified SAPS (>99% pure as judged by HPLC) was supplied as a gift from Vaccine Technology (Patent Cooperation Treaty publication no. WO 2008/068621), having been synthesized on their behalf by Avanti Polar Lipids. alamarBlue was obtained from BioSource International. Cytochalasin D and staurosporine were from Sigma-Aldrich and Calbiochem (Merck Chemicals), respectively. Nitrocellulose membrane and ECL reagent for Western blotting were from GE Healthcare. Anti-phosho-ERK1/2, anti-phospho-JNK, anti-phospho-IκBα, and anti-rabbit secondary Abs were from Cell Signaling Technology. Anti-phospho-p38 was from Promega. Anti-actin was from Sigma-Aldrich. For flow cytometry, PE-conjugated anti-TLR4 mAb (clone HTA125, isotype IgG2a) and PE-conjugated CD14 mAb (clone 61D3, isotype IgG1) and isotype controls were from eBioscience. For fluorescence resonance energy transfer (FRET) experiments, repurified LPS from Salmonella minnesota was purchased from List Biological Laboratories. Lipoteichoic acid (LTA) from S. aureus was a generous gift from Professor Thomas Hartung, University of Konstanz. TLR4- and TLR2-specific mAbs, HTA125 and TL2.3, were purchased from Hycult Biotechnology, while the TLR6-specific polyclonal Ab was from Autogen Bioclear. The CD14-specific mAb MY4 was from BioGenex. Hybridoma cells secreting 26ic (anti-CD14), and W6/32 secreting MHC class I-specific mAbs were from American Type Culture Collection (ATCC). Cholera toxin was purchased from List Biological Laboratories. The Abs used for FRET studies were conjugated to either Cy3 or Cy5 using labeling kits from Amersham Biosciences.

PBMCs were prepared from the venous blood of healthy volunteers taken with informed consent in accordance with a protocol approved by South Sheffield Local Research Ethics Committee. PBMCs were enriched by centrifugation over density gradients as described (27, 28). Monocytes were enriched further by negative magnetic selection using Monocyte Isolation Kit II (Miltenyi Biotec) to a typical mean (±SEM) purity of 83.75 ± 4% (n = 4). The immortalized bronchial epithelial cell line BEAS-2B was maintained in RPMI 1640 containing 2 mM l-glutamine, supplemented with 10% FCS, 100 U/ml penicillin, and 100 μg/ml streptomycin. The monocytic cell line THP-1 was obtained from the ATCC and maintained in RPMI 1640 (supplemented as BEAS-2B). Cells were differentiated in 24-well plates with 0.5 μM PMA (Sigma-Aldrich, Poole, U.K.) for 3 h, nonadherent cells were discarded, and the adherent cells were cultured in normal media for 24 h before use. HUVECs were isolated from umbilical cords, which were donated with informed consent following a protocol approved by North Sheffield Local Research Ethics Committee. Cells were maintained in RPMI 1640 media supplemented with 2 mM l-glutamine, 20 μg/ml endothelial cell growth supplement (Harbor Bio-Products), 95 μg/ml heparin, 0.225% sodium bicarbonate, 100 U/ml penicillin, 100 μg/ml streptomycin, 10% FCS, and 10% newborn calf serum.

Cocultures of BEAS-2B epithelial cells and PBMCs were created (in 24-well plates) with the addition of 30,000 PBMCs/well to 80–90% confluent BEAS-2B cells (seeded 24–48 h before use to attain this confluency), giving a ratio of approximately 1 PBMC to 3 BEAS-2B cells. Monoculture controls were included in all experiments. HUVECs were used at passage 2–3 for all experiments. Cells were seeded in 12-well tissue culture plates and grown to 70–90% confluence, then washed and media replaced with low serum media (2%) for 24 h. HUVEC/monocyte cocultures were created through the addition of 20,000 enriched monocytes/well to the HUVEC monolayers, giving a ratio of ∼1 monocyte to 5 HUVECs. Monoculture controls were included in all experiments.

Cell treatment occurred in the same media as maintenance (see above); in brief, cells were pretreated with SAPS for 1 h before the addition of the TLR or cytokine agonists for 24 h (unless otherwise stated), with SAPS remaining present throughout. Each experiment was conducted multiple times using separate PBMC donors (19 donors used in total) and BEAS-2B or HUVEC cell culture passages.

Cell-free supernatants were collected and stored at −80°C until use. Immunoreactive CXCL8, IL-1β, IL-6, and IL-10 were quantified by ELISA using matched Ab pairs from R&D Systems. The detection limits were 62.5, 19.5, 78, and 32.5 pg/ml respectively. CXCL10 was quantified using a CBA human soluble protein flex set (BD Biosciences) and a FACSArray bioanalyzer (BD Biosciences), in accordance with the manufacturer’s protocols. The limit of detection was 10 pg/ml. Samples whose cytokine levels were undetectable were assigned the detection limit values for graphing and analysis.

Western blot analysis was conducted as described (29). The Abs used were anti-phospho-ERK1/2 (1/500), anti-phospho-JNK (1/500), anti-phospho p38 (1/1000), anti-phospho-IκBα (1/1000), or anti-actin (1/10,000), all detected using a HRP-coupled anti-rabbit secondary Ab (1/2000) followed by enhanced chemiluminescense. Films were densitometrically analyzed using NIH Image (version 1.62f).

Staining and flow cytometry were conducted as previously described (30). Briefly, cells were washed in ice-cold FACS buffer (10 mM PBS without Ca2+ and Mg2+, containing 10 mM HEPES and 0.25% BSA) before nonspecific binding was blocked with mouse IgG (50 μg/ml) and cells were stained with anti-TLR4 or anti-CD14 (both a 1/25 dilution), or isotype control for 45 min at 4°C. Excess Ab was removed by washing, cells were fixed in 1× CellFix (BD Biosciences), and cytometry was performed on a dual laser FACSCalibur using CellQuest software (BD Biosciences). Data were analyzed using FlowJo software version 8.5.3 (Tree Star).

BEAS-2B cells or 300,000 PBMCs were seeded in 96-well plates as described above. Triplicate wells were pretreated with SAPS for 1 h before the addition of 1 ng/ml LPS for 24 h, with SAPS remaining present throughout. Positive controls to induce cell death in BEAS-2B and PBMCs were cytochalasin D (5 μg/ml) and staurosporine (1 μM), respectively. At 20 h, 10% alamarBlue was added and the cells were incubated for a further 4 h. Fluorescence was measured on an fMax fluorometer and Softmax Pro software (Molecular Devices) with excitation at 544 nm and emission at 590 nm. The mean of the absolute fluorescence units for the triplicate wells was calculated, minus the average fluorescence units of media alone (no cells), and data are expressed as percentage change from control cells.

FRET was conducted as described previously (31, 32). Briefly, monocytes were isolated from human A+ buffy coats and cultured on microchamber culture slides (Lab-Tek, Invitrogen) in serum-free media supplemented with 0.01% l-glutamine and 40 μg/ml gentamicin. Cells were labeled with 100 μl of a 1/1 mixture of donor-conjugated Ab (Cy3) and acceptor-conjugated Ab (Cy5). The cells were pretreated with buffer or SAPS (100 μg/ml) for 1 h before stimulation with LPS from S. minnesota (100 ng/ml), or LTA from S. aureus (10 μg/ml), for 10 min. The cells were washed twice in PBS/0.02% BSA before fixation with 4% formaldehyde for 15 min to prevent potential reorganization of the proteins during the course of the experiment. Cells were imaged on a Carl Zeiss LSM510 META confocal microscope (with an Axiovert 200 fluorescent microscope) using a 1.4 NA ×63 Zeiss objective and images were analyzed using LSM image analysis software (Carl Zeiss). The different fluorophores, Cy3 and Cy5, were detected using the appropriate filter sets. Using typical exposure times for image acquisition (<5 s), no fluorescence was observed from a Cy3-labeled specimen using the Cy5 filters, nor was Cy5 fluorescence detected using the Cy3 filter sets. The rate of energy transfer is inversely proportional to the sixth power of the distance between donor and acceptor. The efficiency of energy transfer (E) is defined with respect to r and R0, the characteristic Forster distance by: E = 1/[1 + (r/R0)6]. Energy transfer was detected as an increase in donor fluorescence (dequenching) after complete photobleaching of the acceptor molecule by: E (%) × 100 = 10,000 × [(Cy3 postbleach − Cy3 prebleach)/Cy3 postbleach]. The scaling factor of 10,000 was used to expand E to the scale of the 12-bit images.

All data are presented as means ± SEM (where appropriate) of at least three independent experiments on separate donors (19 donors used in total) and THP-1, BEAS-2B, or HUVEC cell culture passages. Data were analyzed using the statistical tests stated, with ANOVA and the indicated posttest being used for multiple comparisons. Data were analyzed using Prism (version 4.03, GraphPad).

We have previously reported that coculture of immortalized airway epithelial cells with PBMCs results in a marked potentiation of LPS-induced cytokine release when compared with that of either cell type alone (22). Herein, we show that release of the proinflammatory cytokines IL-1β and CXCL8 from epithelial cell/PBMC cocultures in response to LPS was substantially greater than that seen from either cell type alone, and that this was dose-dependently and potently inhibited by SAPS (Fig. 1, A and D). Effective activation of cocultures by LPS is dependent upon activation of monocytes and their initial production of IL-1β (23). We therefore determined whether the modified PS acted on PBMCs to reduce their ability to activate epithelial cells, or upon the epithelial cells to reduce their sensitivity to activation by PBMCs. SAPS inhibited LPS-induced cytokine release from PBMCs alone (Fig. 1, B and E), while the BEAS-2B cells alone did not release either IL-1β or CXCL8 at detectable levels when stimulated with the concentrations of LPS used here (Fig. 1, C and F). A small increase in CXCL8 release was observed in BEAS-2B cells in response to the higher doses of SAPS (Fig. 1 F), indicating that SAPS may have some modest proinflammatory effects, although these were restricted to tissue cells.

FIGURE 1.

Inhibition of LPS-induced cytokine release from epithelial cell/PBMC cocultures and PBMC monocultures by PS. The airway epithelial cell line BEAS-2B was grown to confluence in 24-well plates, and cocultures were created through the addition of 30,000 PBMCs per well (A and D). Monoculture controls of 30,000 PBMCs (B and E) or BEAS-2B cells (C and F) alone were also created. Cells were treated with SAPS for 1 h at the doses indicated, before addition of media or LPS (0.1 or 1 ng/ml). After 24 h, cell-free supernatants were prepared and levels of IL-1β (A–C) and CXCL8 (D–F) measured by ELISA. Data shown are means ± SEM (n = 5), with each replicate performed at a separate passage with freshly prepared PBMCs from independent donors. Significant differences in cytokine production are indicated by ∗, p < 0.05, ∗∗, p < 0.01, and ∗∗∗, p < 0.001, analyzed by two-way ANOVA and Bonferroni’s posttest.

FIGURE 1.

Inhibition of LPS-induced cytokine release from epithelial cell/PBMC cocultures and PBMC monocultures by PS. The airway epithelial cell line BEAS-2B was grown to confluence in 24-well plates, and cocultures were created through the addition of 30,000 PBMCs per well (A and D). Monoculture controls of 30,000 PBMCs (B and E) or BEAS-2B cells (C and F) alone were also created. Cells were treated with SAPS for 1 h at the doses indicated, before addition of media or LPS (0.1 or 1 ng/ml). After 24 h, cell-free supernatants were prepared and levels of IL-1β (A–C) and CXCL8 (D–F) measured by ELISA. Data shown are means ± SEM (n = 5), with each replicate performed at a separate passage with freshly prepared PBMCs from independent donors. Significant differences in cytokine production are indicated by ∗, p < 0.05, ∗∗, p < 0.01, and ∗∗∗, p < 0.001, analyzed by two-way ANOVA and Bonferroni’s posttest.

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These data suggest that the main mechanism by which our modified PS inhibited inflammation in our model was through down-regulation of proinflammatory effects during the initial PBMC response. However, the antiinflammatory cytokine IL-10 is produced by monocytes and is a potent suppressor of proinflammatory cytokine production, including IL-1β and CXCL8 (33). Consequently, we investigated whether SAPS could directly increase IL-10 production from PBMCs and potentially provide a mechanism for suppression of LPS-induced proinflammatory cytokine release. Our data revealed that SAPS dose-dependently inhibited LPS-induced IL-10 release from PBMCs (Fig. 2), and thus our modified PS appeared to have a global inhibitory effect on TLR4 function.

FIGURE 2.

The inhibitory effects of SAPS are not through enhanced production of the antiinflammatory cytokine IL-10. PBMCs (300,000) were treated with media or SAPS (at the doses indicated) for 1 h before addition of LPS (1 ng/ml) for 24 h. Cell-free supernatants were prepared and levels of IL-10 measured by ELISA. Data shown are means ± SEM, with each replicate performed with freshly prepared PBMCs from three independent donors. Significant differences in cytokine production are indicated by ∗, p < 0.05 and ∗∗, p < 0.01, analyzed by one-way ANOVA and Tukey’s posttest.

FIGURE 2.

The inhibitory effects of SAPS are not through enhanced production of the antiinflammatory cytokine IL-10. PBMCs (300,000) were treated with media or SAPS (at the doses indicated) for 1 h before addition of LPS (1 ng/ml) for 24 h. Cell-free supernatants were prepared and levels of IL-10 measured by ELISA. Data shown are means ± SEM, with each replicate performed with freshly prepared PBMCs from three independent donors. Significant differences in cytokine production are indicated by ∗, p < 0.05 and ∗∗, p < 0.01, analyzed by one-way ANOVA and Tukey’s posttest.

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TLR4 function is, in part, regulated by control of its expression and shuttling of the receptor from endosomal to plasma membrane compartments (34, 35). We therefore determined if exposure to PS, a membrane lipid, exerted effects on TLR surface expression that might underpin inhibition of TLR4 responses. We found that incubation of PBMCs with SAPS for 1 h had little effect on TLR4 or CD14 expression as measured by flow cytometry (Fig. 3, A and B). To rule out the possibility that SAPS was mediating its inhibitory effects by having detrimental actions on the cells, thus diminishing their ability to produce cytokines, we assessed the ability of PBMCs and epithelial cells to reduce the dye alamarBlue as a measure of their metabolic capacity and hence viability (36, 37). The metabolic capacity of PBMCs and epithelial cells was unaffected by SAPS (with or without LPS) (Fig. 3, C and D). These data suggested that SAPS was not acting to globally disturb membrane functions such as uptake of nutrients, was nontoxic, and did not interfere with cellular metabolism.

FIGURE 3.

The actions of SAPS are not through regulation of TLR4/CD14 surface expression or due to cell death. Cell surface expression of TLR4 (A) and CD14 (B) on PBMCs was measured by flow cytometric analysis after treatment with media or SAPS (50 μg/ml) for 1 h, before staining the cells with the anti-TLR4, anti-CD14 mAbs, or isotype controls. Representative histograms are shown (n = 4 PBMCs from separate donors). Viability of PBMCs (C) and BEAS-2Bs (D) were measured by reduction of alamarBlue. PBMCs (300,000) or BEAS-2B (300,000) cells were seeded in 96-well plates and triplicate wells were treated with SAPS for 1 h before the addition of 1 ng/ml LPS for 24 h, with SAPS remaining present throughout. At 20 h, 10% alamarBlue was added and the cells incubated for a further 4 h. Positive controls to induce cell death were staurosporine (1 μM) for PBMCs and cytochalasin D (5 μg/ml) for BEAS-2B. The mean of the absolute fluorescence units for the triplicate wells was calculated, minus the average fluorescence units of media alone (no cells), and data are expressed as percentage change from control cells. Data shown are means ± SEM of n = 4 for PBMCs and n = 3 for BEAS-2B cells, with each replicate performed on independent donors or at a separate passage, respectively.

FIGURE 3.

The actions of SAPS are not through regulation of TLR4/CD14 surface expression or due to cell death. Cell surface expression of TLR4 (A) and CD14 (B) on PBMCs was measured by flow cytometric analysis after treatment with media or SAPS (50 μg/ml) for 1 h, before staining the cells with the anti-TLR4, anti-CD14 mAbs, or isotype controls. Representative histograms are shown (n = 4 PBMCs from separate donors). Viability of PBMCs (C) and BEAS-2Bs (D) were measured by reduction of alamarBlue. PBMCs (300,000) or BEAS-2B (300,000) cells were seeded in 96-well plates and triplicate wells were treated with SAPS for 1 h before the addition of 1 ng/ml LPS for 24 h, with SAPS remaining present throughout. At 20 h, 10% alamarBlue was added and the cells incubated for a further 4 h. Positive controls to induce cell death were staurosporine (1 μM) for PBMCs and cytochalasin D (5 μg/ml) for BEAS-2B. The mean of the absolute fluorescence units for the triplicate wells was calculated, minus the average fluorescence units of media alone (no cells), and data are expressed as percentage change from control cells. Data shown are means ± SEM of n = 4 for PBMCs and n = 3 for BEAS-2B cells, with each replicate performed on independent donors or at a separate passage, respectively.

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LPS activates the TLR4 receptor complex triggering two downstream signaling pathways termed MyD88-dependent and -independent, governed by the adaptor proteins that are recruited (38).

Production of proinflammatory cytokines, including IL-1β and CXCL8, is heavily dependent on the MyD88-dependent arm that subsequently leads to activation of early-phase NF-κB and the MAP kinases, which diverge at the level of the TRAF6/TAB2/TAK1/TAB1 complex (38). To gain further insights into the level at which SAPS exerted its actions, we investigated the actions of SAPS on LPS-mediated NF-κB and MAP kinase signaling. Treatment of PBMCs with LPS for 30 min activated NF-κB signaling as shown by IκBα phosphorylation, and this was inhibited by a 1-h pretreatment with SAPS (Fig. 4,A). SAPS also inhibited phosphorylation of the MAP kinase family members JNK (Fig. 4,B), p38 (Fig. 4,C), and ERK (Fig. 4 D).

FIGURE 4.

Inhibition of LPS signaling involves inhibition of the MyD88-dependent signaling pathway. PBMCs (300,000) were treated with media or SAPS (10 or 50 μg/ml) for 1 h before addition of LPS (1 ng/ml) for 30 min. Whole-cell lysates were analyzed by Western blot using Abs specific to the phosphorylated forms of IκBα (A), JNK (p46/54) (B), p38 (C), or ERK (p42/44) (D). Blots were densitometrically analyzed and normalized to β-actin to control for loading. Data are presented as means ± SEM of n = 4 (A and C) or n = 3 (B and D) separate experiments from independent donors with representative Western blots shown below. Significant differences are indicated by ∗, p < 0.05, ∗∗, p < 0.01, and ∗∗∗, p < 0.001, analyzed by one-way ANOVA and Tukey’s posttest.

FIGURE 4.

Inhibition of LPS signaling involves inhibition of the MyD88-dependent signaling pathway. PBMCs (300,000) were treated with media or SAPS (10 or 50 μg/ml) for 1 h before addition of LPS (1 ng/ml) for 30 min. Whole-cell lysates were analyzed by Western blot using Abs specific to the phosphorylated forms of IκBα (A), JNK (p46/54) (B), p38 (C), or ERK (p42/44) (D). Blots were densitometrically analyzed and normalized to β-actin to control for loading. Data are presented as means ± SEM of n = 4 (A and C) or n = 3 (B and D) separate experiments from independent donors with representative Western blots shown below. Significant differences are indicated by ∗, p < 0.05, ∗∗, p < 0.01, and ∗∗∗, p < 0.001, analyzed by one-way ANOVA and Tukey’s posttest.

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The MyD88-independent pathway involves recruitment of the adaptors TRIF (TIR domain-containing adaptor-inducing IFN-β) and TRAM (TRIF-related adaptor molecule), leading to activation of late-phase NF-κB and a second pathway involving activation of the transcription factor IFN response factor 3 (IRF-3) and production of IFN-inducible cytokines such as IFN-inducible protein 10 (IP-10/CXCL10) (39). We investigated activation of this pathway in a macrophage-like cell line, differentiated THP-1 cells. Treatment of THP-1 cells with a low (10 ng/ml) and high (1 μg/ml) dose of LPS for 24 h resulted in release of CXCL10 (Fig. 5,A) and IL-1β (Fig. 5 B; used to confirm SAPS was an effective inhibitor of LPS responses in THP-1 cells), which was significantly inhibited in the presence of SAPS. These data show SAPS potently inhibits both arms of the TLR4 signaling pathway and indicates it is most likely mediating its effects at the level of the receptor and/or adaptor complex.

FIGURE 5.

The LPS-induced MyD88-independent signaling pathway is inhibited by SAPS. Differentiated THP-1 cells were treated with SAPS (50 μg/ml) for 1 h before the addition of LPS (10 or 1000 ng/ml) for 24 h, with SAPS remaining present throughout. Cell-free supernatants were prepared and levels of CXCL10 (A) or IL-1β (B) measured by CBA or ELISA, respectively. Data are means ± SEM of three experiments, with each performed on a separate cell passage. Significant differences are indicated by ∗, p < 0.05 and ∗∗∗, p < 0.001, analyzed by one-way ANOVA and Tukey’s posttest.

FIGURE 5.

The LPS-induced MyD88-independent signaling pathway is inhibited by SAPS. Differentiated THP-1 cells were treated with SAPS (50 μg/ml) for 1 h before the addition of LPS (10 or 1000 ng/ml) for 24 h, with SAPS remaining present throughout. Cell-free supernatants were prepared and levels of CXCL10 (A) or IL-1β (B) measured by CBA or ELISA, respectively. Data are means ± SEM of three experiments, with each performed on a separate cell passage. Significant differences are indicated by ∗, p < 0.05 and ∗∗∗, p < 0.001, analyzed by one-way ANOVA and Tukey’s posttest.

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The proinflammatory cytokine IL-1β also activates intracellular signaling via MyD88. Consequently, were SAPS to be acting at the level of the adaptor complex, it would likely prevent cellular responses to IL-1β. However, SAPS had no effect on IL-1β-induced CXCL8 release from cocultures (Fig. 6,A) or on the control monocultures (Fig. 6, B and C). These data demonstrated that inhibition of LPS signaling was likely to be specific, was not mediated through nonselective disruption of cytokine processing or release, and was occurring at the level of the TLR4 complex.

FIGURE 6.

IL-1β-induced cytokine release is unaffected by membrane microdomain disruption. BEAS-2B epithelial cells were grown to confluence in 24-well plates, and cocultures were created through the addition of 30,000 PBMCs per well (A and D). Monoculture controls of 30,000 PBMCs (B) and BEAS-2B cells (C) were also created. Cells were treated with SAPS for 1 h at the doses indicated, before addition of media, IL-1β (0.1 or 1 ng/ml), LPS (0.1 ng/ml), or LPS (0.1 ng/ml) + IL-1β (1 ng/ml) costimulation. After 24 h, cell-free supernatants were prepared and levels of CXCL8 measured by ELISA. Data shown are mean ± SEMs of n = 5 (A–C) or n = 3 (D), with each replicate performed at a separate passage with freshly prepared PBMCs from independent donors. Data were analyzed by two-way ANOVA and Bonferroni’s posttest.

FIGURE 6.

IL-1β-induced cytokine release is unaffected by membrane microdomain disruption. BEAS-2B epithelial cells were grown to confluence in 24-well plates, and cocultures were created through the addition of 30,000 PBMCs per well (A and D). Monoculture controls of 30,000 PBMCs (B) and BEAS-2B cells (C) were also created. Cells were treated with SAPS for 1 h at the doses indicated, before addition of media, IL-1β (0.1 or 1 ng/ml), LPS (0.1 ng/ml), or LPS (0.1 ng/ml) + IL-1β (1 ng/ml) costimulation. After 24 h, cell-free supernatants were prepared and levels of CXCL8 measured by ELISA. Data shown are mean ± SEMs of n = 5 (A–C) or n = 3 (D), with each replicate performed at a separate passage with freshly prepared PBMCs from independent donors. Data were analyzed by two-way ANOVA and Bonferroni’s posttest.

Close modal

We examined whether addition of exogenous IL-1β could restore LPS-induced coculture responses in the presence of SAPS. We found that dual stimulation of our coculture model with LPS and IL-1β produced an additive effect on CXCL8 release (Fig. 6,D). Dual stimulation with exogenous IL-1β rendered SAPS a less effective inhibitor of coculture activation, suggesting that IL-1β could partially restore CXCL8 release. However at 50 μg/ml, SAPS completely inhibited LPS responses and CXCL8 release was reduced to the level produced by IL-1β alone (Fig. 6 D).

LPS stimulation results in segregation of TLR4 into membrane microdomains (7, 8). We hypothesized that exposure of cells to SAPS would result in altered membrane biology and disturb microdomain function. We investigated this possibility using a FRET-based approach to determine consequences of SAPS on TLR4/CD14 association. FRET was measured in terms of dequenching of donor fluorescence after complete photobleaching of the acceptor fluorophore. Increased donor fluorescence, after complete destruction of the acceptor, indicated that the donor fluorescence was quenched in the presence of the acceptor because of energy transfer. The energy transfer efficiency of the system was tested using, as a positive control, the energy transfer from Cy3-26ic and Cy5-MY4 (mAbs to two different epitopes on CD14), which showed that the maximum energy transfer efficiency (E) was 37 ± 1.5% in unstimulated monocytes, which was not affected by treatment with SAPS or LPS, alone or in combination (Table I). A negative control comprising Cy3-W6/32 (the mAb specific for MHC class I) and Cy5-cholera toxin (which recognizes GM-1 ganglioside; a raft-associated lipid) was also used, which revealed no significant energy transfer in any of our four experimental groups (Table I).

Table I.

SAPS disrupts the association of TLR4 and its receptor partners within membrane microdomainsa

Donor (Cy3)Acceptor (Cy5)TreatmentE ± ΔE (%)
CD14 CD14 Control 37 ± 1.5  
  SAPS 38 ± 0.2  
  LPS 36 ± 2.0  
  LPS + SAPS 38 ± 0.5  
MHC class I GM1 Control 5 ± 1.5  
  SAPS 6 ± 1.0  
  LPS 6 ± 1.0  
  LPS + SAPS 6 ± 0.5  
CD14 GM1 Control 36 ± 1.0] ∗∗∗ 
  SAPS 9 ± 1.5  
  LPS 35 ± 0.5] ∗∗∗ 
  LPS + SAPS 10 ± 0.6  
TLR4 GM1 Control 6 ± 0.5  
  SAPS 7 ± 1.5  
  LPS 37 ± 1.5] ∗∗∗ 
  LPS + SAPS 17 ± 1.0  
TLR4 CD14 Control 7 ± 1.0  
  SAPS 8 ± 0.8  
  LPS 32 ± 0.5] ∗∗∗ 
  LPS + SAPS 12 ± 1.5  
Donor (Cy3)Acceptor (Cy5)TreatmentE ± ΔE (%)
CD14 CD14 Control 37 ± 1.5  
  SAPS 38 ± 0.2  
  LPS 36 ± 2.0  
  LPS + SAPS 38 ± 0.5  
MHC class I GM1 Control 5 ± 1.5  
  SAPS 6 ± 1.0  
  LPS 6 ± 1.0  
  LPS + SAPS 6 ± 0.5  
CD14 GM1 Control 36 ± 1.0] ∗∗∗ 
  SAPS 9 ± 1.5  
  LPS 35 ± 0.5] ∗∗∗ 
  LPS + SAPS 10 ± 0.6  
TLR4 GM1 Control 6 ± 0.5  
  SAPS 7 ± 1.5  
  LPS 37 ± 1.5] ∗∗∗ 
  LPS + SAPS 17 ± 1.0  
TLR4 CD14 Control 7 ± 1.0  
  SAPS 8 ± 0.8  
  LPS 32 ± 0.5] ∗∗∗ 
  LPS + SAPS 12 ± 1.5  
a

Energy transfer between different pairs was detected from the increase in donor fluorescence after acceptor photobleaching. Data represent means ± SD of three independent experiments. Significant effects of SAPS are represented by ∗∗∗, p < 0.001; analyzed by one-way ANOVA and Tukey’s posttest.

Previously studies have shown that CD14 is situated within membrane microdomains in a resting state, while TLR4 moves into these domains only upon stimulation with LPS (32), and that LPS triggers a physical association between CD14 and TLR4 (40). In this study we confirmed these findings and reveal that SAPS disrupts this association. The results revealed large dequenching between CD14/GM-1 ganglioside in control cells confirming CD14 resides in membrane microdomains and treatment with LPS has no effect while SAPS disrupt this association both in the presence and absence of LPS (Table I). As expected there is no association between TLR4 and either GM-1 ganglioside or CD14 in the resting cell, but upon LPS stimulation a large dequenching occurs between TLR4/GM-1 and TLR4/CD14, revealing that TLR4 concentrates in microdomains after LPS stimulation and associates with CD14 within the membrane. Finally, monocytes stimulated with LPS in the presence of SAPS show reduced energy transfer in both cases, indicating that SAPS causes disassembly of these microdomains and a consequent inability of TLR4 and CD14 to associate (Table I).

A likely role for membrane microdomains in TLR4 signaling is established, but the importance of these domains in signaling of other TLRs is less well explored, and the potential for membrane disruption to affect highly geared inflammatory models such as our coculture systems is equally uncertain. We therefore explored the consequences of SAPS on inflammatory responses induced by a range of TLR and non-TLR agonists. We selected TLR agonists signaling from the cell surface and from endosomes to gain insights into the dependence of TLR signaling on membrane microdomains.

We first determined the extent to which maximal epithelial cell responses to TLR and non-TLR agonists depended on cooperative signaling. We stimulated cocultures of BEAS-2B and PBMCs, and their monoculture controls, with Pam3CSK4 (TLR2/1), Gardiquimod (which signals from acidified endosomes, activating TLR7 > TLR8), and the proinflammatory cytokines TNF-α and IL-1β (Fig. 7). Epithelial cells alone did not respond to Pam3CSK4 or Gardiquimod, although a response to the cytokines TNF-α and particularly IL-1β was observed. In contrast, PBMCs showed a modest response to the TLR agonists but not the cytokines. However, when the same numbers of cells were cultured together a significantly enhanced response to all four agonists was observed, demonstrating that tissue cells and infiltrating leukocytes will interact and cooperate in response to both exogenous pathogens and endogenous proinflammatory cytokines.

FIGURE 7.

PBMCs enhance epithelial responses to TLR and non-TLR agonists. BEAS-2B epithelial cells were grown to confluence in 24-well plates, and cocultures were created through the addition of 30,000 PBMCs per well. Monoculture controls of 30,000 PBMCs and BEAS-2B cells were also created. Cells were treated with media or Pam3CSK4, Gardiquimod, TNF-α, or IL-1β at the doses indicated. After 24 h, cell-free supernatants were prepared and levels of CXCL8 measured by ELISA. Data shown are means ± SEM of n = 5, with each replicate performed at a separate passage with freshly prepared PBMCs from independent donors. Significant differences in cytokine production between coculture and BEAS-2B cells or PBMCs are indicated by ∗∗, p < 0.01, ∗∗∗, p < 0.001, or +, p < 0.05, +++, p < 0.001, respectively, analyzed by two-way ANOVA and Bonferroni’s posttest.

FIGURE 7.

PBMCs enhance epithelial responses to TLR and non-TLR agonists. BEAS-2B epithelial cells were grown to confluence in 24-well plates, and cocultures were created through the addition of 30,000 PBMCs per well. Monoculture controls of 30,000 PBMCs and BEAS-2B cells were also created. Cells were treated with media or Pam3CSK4, Gardiquimod, TNF-α, or IL-1β at the doses indicated. After 24 h, cell-free supernatants were prepared and levels of CXCL8 measured by ELISA. Data shown are means ± SEM of n = 5, with each replicate performed at a separate passage with freshly prepared PBMCs from independent donors. Significant differences in cytokine production between coculture and BEAS-2B cells or PBMCs are indicated by ∗∗, p < 0.01, ∗∗∗, p < 0.001, or +, p < 0.05, +++, p < 0.001, respectively, analyzed by two-way ANOVA and Bonferroni’s posttest.

Close modal

We then investigated the potential for such systems to be disrupted by SAPS. There is some evidence for a role for CD14 in TLR2 signaling (41, 42, 43), and we therefore explored whether SAPS would disrupt TLR2 responses. We found that SAPS dose-dependently and significantly inhibited IL-1β release from epithelial cell/PBMC cocultures stimulated with Pam3CSK4 (Fig. 8 A). As with responses to TLR4, the ability of SAPS to inhibit activation of cocultures was again likely due to actions on the PBMCs, since SAPS also inhibited Pam3CSK4-induced IL-1β release from PBMCs monoculture controls, while the BEAS-2B cells alone showed no response to Pam3CSK4 (data not shown). Similar results were obtained for Pam3CSK4-induced CXCL8 release (data not shown).

FIGURE 8.

Membrane microdomain disruption inhibits Pam3CSK4- and S. pneumoniae-induced cytokine release. BEAS-2B epithelial cells were grown to confluence in 24-well plates, and cocultures were created through the addition of 30,000 PBMCs per well. Cells were treated with SAPS for 1 h at the doses indicated, before addition of Pam3CSK4 at the doses indicated (A). Alternately, 300,000 PBMCs were treated with media or SAPS (at the indicated doses) for 1 h before addition of heat killed S. pneumoniae (at a ratio of 1:1, 5:1, and 10:1 S. pneumoniae/PBMCs) (B). After 24 h, cell-free supernatants were prepared and levels of IL-1β measured by ELISA. Data shown are meana ± SEM of n = 4 (A) or n = 5 (B), with each replicate performed at a separate cell passage and/or with freshly prepared PBMCs from independent donors. Significant differences in cytokine production are indicated by ∗, p < 0.05, ∗∗, p < 0.01, and ∗∗∗, p < 0.001, analyzed by two-way ANOVA and Bonferroni’s posttest.

FIGURE 8.

Membrane microdomain disruption inhibits Pam3CSK4- and S. pneumoniae-induced cytokine release. BEAS-2B epithelial cells were grown to confluence in 24-well plates, and cocultures were created through the addition of 30,000 PBMCs per well. Cells were treated with SAPS for 1 h at the doses indicated, before addition of Pam3CSK4 at the doses indicated (A). Alternately, 300,000 PBMCs were treated with media or SAPS (at the indicated doses) for 1 h before addition of heat killed S. pneumoniae (at a ratio of 1:1, 5:1, and 10:1 S. pneumoniae/PBMCs) (B). After 24 h, cell-free supernatants were prepared and levels of IL-1β measured by ELISA. Data shown are meana ± SEM of n = 4 (A) or n = 5 (B), with each replicate performed at a separate cell passage and/or with freshly prepared PBMCs from independent donors. Significant differences in cytokine production are indicated by ∗, p < 0.05, ∗∗, p < 0.01, and ∗∗∗, p < 0.001, analyzed by two-way ANOVA and Bonferroni’s posttest.

Close modal

We then determined whether SAPS disrupted the ability of TLR2 to associate with its signaling partners in membrane microdomains. A large dequenching was observed between TLR4/CD14 after LPS (Table I and II), but not after LTA (Table II), stimulation, and SAPS disrupted this association (Table II). In contrast, TLR2/CD14 associated after LTA, but not LPS, stimulation, and treatment with SAPS prevented this association (Table II). Treatment of monocytes with LTA also resulted in association of TLR2/GM-1 ganglioside and TLR6/GM-1 ganglioside, revealing that these receptors both concentrated within the microdomains of activated cells. These associations were disrupted by treatment of cells with SAPS (Table II).

Table II.

SAPS disrupts the association of TLR2 and its receptor partners within membrane microdomainsa

Donor (Cy3)Acceptor (Cy5)TreatmentE ± ΔE (%)
TLR4 CD14 Control 6 ± 1.0  
  SAPS 5 ± 0.5  
  LPS 34 ± 1.5] ∗∗∗ 
  LPS + SAPS 13 ± 1.5  
  LTA 6 ± 0.5  
  LTA + SAPS 7 ± 0.8  
TLR2 CD14 Control 6 ± 1.0  
  SAPS 7 ± 1.2  
  LPS 5 ± 0.5  
  LPS + SAPS 6 ± 0.4  
  LTA 35 ± 0.8] ∗∗∗ 
  LTA + SAPS 12 ± 0.8  
TLR2 GM1 Control 7 ± 0.8  
  SAPS 6 ± 1.0  
  LPS 7 ± 1.0  
  LPS + SAPS 8 ± 1.0  
  LTA 36 ± 1.0] ∗∗∗ 
  LTA + SAPS 14 ± 1.0  
TLR6 GM1 Control 7 ± 0.8  
  SAPS 6 ± 0.5  
  LPS 7 ± 1.0  
  LPS + SAPS 5 ± 0.8  
  LTA 35 ± 1.0] ∗∗∗ 
  LTA + SAPS 13 ± 1.2  
Donor (Cy3)Acceptor (Cy5)TreatmentE ± ΔE (%)
TLR4 CD14 Control 6 ± 1.0  
  SAPS 5 ± 0.5  
  LPS 34 ± 1.5] ∗∗∗ 
  LPS + SAPS 13 ± 1.5  
  LTA 6 ± 0.5  
  LTA + SAPS 7 ± 0.8  
TLR2 CD14 Control 6 ± 1.0  
  SAPS 7 ± 1.2  
  LPS 5 ± 0.5  
  LPS + SAPS 6 ± 0.4  
  LTA 35 ± 0.8] ∗∗∗ 
  LTA + SAPS 12 ± 0.8  
TLR2 GM1 Control 7 ± 0.8  
  SAPS 6 ± 1.0  
  LPS 7 ± 1.0  
  LPS + SAPS 8 ± 1.0  
  LTA 36 ± 1.0] ∗∗∗ 
  LTA + SAPS 14 ± 1.0  
TLR6 GM1 Control 7 ± 0.8  
  SAPS 6 ± 0.5  
  LPS 7 ± 1.0  
  LPS + SAPS 5 ± 0.8  
  LTA 35 ± 1.0] ∗∗∗ 
  LTA + SAPS 13 ± 1.2  
a

Energy transfer between different pairs was detected from the increase in donor fluorescence after acceptor photobleaching. Data represent means ± SD of three independent experiments. Significant effects of SAPS are represented by ∗∗∗, p < 0.001; analyzed by one-way ANOVA and Tukey’s posttest.

Finally, S. pneumoniae is a leading cause of invasive bacterial disease capable of activating many TLRs and other pattern recognition systems. Notably, SAPS is also capable of significantly inhibiting heat-killed S. pneumoniae-induced IL-1β release from PBMCs (Fig. 8 B). Similar results were obtained for S. pneumoniae-induced CXCL8 release (data not shown).

We subsequently determined if SAPS also disrupted signaling from intracellular compartments in tissue cells. In contrast to TLR2, TLR7/8 is exclusively expressed intracellularly and signals from acidified endosomes (44, 45). Epithelial cell/PBMC coculture responses to a TLR7/8 agonist, Gardiquimod, were also dose-dependently and significantly inhibited by SAPS (Fig. 9,A), which again was due to actions on the PBMCs (data not shown). As TLR3 is expressed by tissue cells but not PBMCs (22), we studied the effects of SAPS on poly(I:C) (a synthetic double-stranded RNA analog) induced CXCL8 release from epithelial cell monocultures. Only the highest dose of SAPS (50 μg/ml) reduced CXCL8 release (Fig. 9,B). The control compound poly(dI:dC) did not cause cytokine release, except in combination with the highest dose of SAPS, which is in keeping with the modest proinflammatory effects of SAPS alone that we observed previously (Fig. 1 F).

FIGURE 9.

Membrane microdomain disruption inhibits Gardiquimod (TLR7/8)-induced cytokine release, but has only a modest effect on poly(I:C) (TLR3)-induced cytokine release. BEAS-2B epithelial cells were grown to confluence in 24-well plates, and cocultures were created through the addition of 30,000 PBMCs per well (A). Monocultures of BEAS-2B cells alone were also created (B). Cells were treated with SAPS for 1 h at the doses indicated, before addition of Gardiquimod (A) or poly(I:C) (B) at the doses indicated. After 24 h, cell-free supernatants were prepared and levels of IL-1β (A) or CXCL8 (B) measured by ELISA. Data shown are means ± SEM of n = 3 (A) or n = 4 (B), with each replicate performed on freshly prepared PBMCs from independent donors and/or at a separate cell passage. Significant differences in cytokine production are indicated by ∗, p < 0.05 and ∗∗∗, p < 0.001, analyzed by two-way ANOVA and Bonferroni’s posttest.

FIGURE 9.

Membrane microdomain disruption inhibits Gardiquimod (TLR7/8)-induced cytokine release, but has only a modest effect on poly(I:C) (TLR3)-induced cytokine release. BEAS-2B epithelial cells were grown to confluence in 24-well plates, and cocultures were created through the addition of 30,000 PBMCs per well (A). Monocultures of BEAS-2B cells alone were also created (B). Cells were treated with SAPS for 1 h at the doses indicated, before addition of Gardiquimod (A) or poly(I:C) (B) at the doses indicated. After 24 h, cell-free supernatants were prepared and levels of IL-1β (A) or CXCL8 (B) measured by ELISA. Data shown are means ± SEM of n = 3 (A) or n = 4 (B), with each replicate performed on freshly prepared PBMCs from independent donors and/or at a separate cell passage. Significant differences in cytokine production are indicated by ∗, p < 0.05 and ∗∗∗, p < 0.001, analyzed by two-way ANOVA and Bonferroni’s posttest.

Close modal

We subsequently sought to compare the effects of SAPS as a potent disruptor of membrane microdomains in a further model system, using our established model of vascular inflammation in which HUVECs are cultured with purified monocytes (22). In this study we showed that IL-6 release from LPS-activated HUVEC/monocyte cocultures was dose-dependently inhibited by SAPS (Fig. 10,A). The inhibitory effect of SAPS in cocultures appears to be through actions on both cell types since SAPS also inhibited LPS-induced IL-6 release from monocytes (Fig. 10,B) and HUVECs (Fig. 10 C) alone. Similar results were obtained for inhibition of LPS-induced IL-1β and CXCL8 release by SAPS (data not shown).

FIGURE 10.

Inhibition of LPS-induced cytokine release in models of vascular inflammation. HUVECs were grown to confluence in 12-well plates and cocultures created through the addition of 20,000 monocytes (enriched from PBMCs by negative magnetic selection) per well (A). Monoculture controls of 20,000 monocytes (B) or HUVECs alone (C) were also created. Cells were treated with SAPS for 1 h at the doses indicated, before addition of media or LPS (0.1 or 1 ng/ml). After 24 h, cell-free supernatants were prepared and levels of IL-6 measured by ELISA. Data shown are means ± SEM of n = 4, with each replicate performed on independent HUVEC and monocyte donors. Significant differences in cytokine production are indicated by ∗, p < 0.05, ∗∗, p < 0.01, and ∗∗∗, p < 0.001, analyzed by two-way ANOVA and Bonferroni’s posttest.

FIGURE 10.

Inhibition of LPS-induced cytokine release in models of vascular inflammation. HUVECs were grown to confluence in 12-well plates and cocultures created through the addition of 20,000 monocytes (enriched from PBMCs by negative magnetic selection) per well (A). Monoculture controls of 20,000 monocytes (B) or HUVECs alone (C) were also created. Cells were treated with SAPS for 1 h at the doses indicated, before addition of media or LPS (0.1 or 1 ng/ml). After 24 h, cell-free supernatants were prepared and levels of IL-6 measured by ELISA. Data shown are means ± SEM of n = 4, with each replicate performed on independent HUVEC and monocyte donors. Significant differences in cytokine production are indicated by ∗, p < 0.05, ∗∗, p < 0.01, and ∗∗∗, p < 0.001, analyzed by two-way ANOVA and Bonferroni’s posttest.

Close modal

The work of this manuscript describes a number of important observations. We provide further evidence for the crucial role of cooperative networks in the mounting of effective inflammatory responses to TLR agonists. We identify a novel, nontoxic inhibitor of TLR responses that exerts broad-ranging inhibition of TLR, but not IL-1β, signaling, and show that this is mediated via disruption of membrane microdomains, extending our knowledge of the potential role of these domains in TLR signaling, including in TLRs located in and signaling from the endosome. Disruption of membrane function nonetheless does not induce cell death or impair cytokine release, demonstrating that microdomain disruption has the potential to be a nontoxic strategy to intervene in inflammatory signaling.

Most in vitro studies of TLR responses consider actions of TLRs on single cell types. In recent publications we have shown that signaling to several TLR agonists is most effectively mediated by cooperative responses (22, 23). In particular, we have defined cooperative networks in which leukocytes, principally the monocyte, can use tissue cells to amplify inflammatory responses, and in which engagement of the tissue cell in the innate immune response is dependent upon such networks. In these systems, early production of IL-1 from the monocyte appears essential for effective tissue cell activation (23). Such networks enable effective tissue responses to agonists to which the tissue cells may be markedly less responsive.

Illustrating these concepts, we herein show that epithelial cells are unresponsive to a TLR7/8 agonist, consistent with the lack of expression of this receptor seen in other epithelial cell lines (46). However, in the presence of PBMCs a profound response to the TLR7/8 agonist is observed as defined by production of proinflammatory cytokines. These data are similar to responses we have previously observed in airway smooth muscle (22), and reinforce the generic dependence of innate immunity on such networks. Even where tissue cells are responsive to TLR agonists such as LPS, we show again here (as previously, see Ref. 23) that signaling is profoundly greater when cooperative networks are established, whether the tissue cells are epithelial or endothelial in origin. Evidence for the in vivo relevance of such systems is accumulating (47).

Despite the high gearing of these networks, they are nonetheless amenable to therapeutic targeting. We have shown previously that IL-1 appears to play a crucial role in initiating inflammatory responses (23). Herein, we identify another way to manipulate these systems, yielding important insights into TLR function and the role of membrane microdomains in cell signaling.

We hypothesized initially that PS species would inhibit LPS-mediated activation of cocultures through antagonism of LPS binding to CD14. We explored this hypothesis using a specific PS species, SAPS, which is found in plasma in vivo (48). In keeping with our first hypothesis, we found that SAPS inhibited TLR4-mediated coculture activation and responses of individual cells, including freshly isolated human PBMCs, differentiated THP-1 cells (a macrophage model), and endothelial cells. In contrast, SAPS did not inhibit the actions of IL-1β in cocultures of epithelial cells and PBMCs, or in either cell type alone. Furthermore, we found that the addition of exogenous IL-1β could partially restore the inflammatory response induced by LPS even in the presence of SAPS, although this was evident only at lower SAPS concentrations (1 and 10 μg/ml). This suggests that while IL-1β is a crucial communicator between leukocytes and tissue cells, other factors are also required to convey full activation of the LPS-induced inflammatory response, and at higher doses of SAPS (50 μg/ml), production of these additional factors is abolished.

We then sought to identify the mechanism of action whereby SAPS mediated its inhibitory effects on proinflammatory cytokine release. We found that this was not through production of the antiinflammatory cytokine IL-10, nor did SAPS cause down-regulation of expression of the TLR4 signaling complex or death of either tissue cells or PBMCs. Although TLR2 signaling may also show some CD14 dependence (41, 42, 43), we were surprised by the degree to which SAPS abolished this. Our finding that SAPS inhibited both the MyD88-dependent and -independent signaling pathways equally strongly suggested that SAPS was acting at the level of the receptor or its immediate signaling complex. However, the lack of actions on IL-1β-induced responses, which also utilize the MyD88 adaptor, suggested actions at the adaptor level were unlikely. We therefore hypothesized that SAPS might be directly influencing membrane dynamics. We found that SAPS disrupted the association of both TLR4 and TLR2 with their respective membrane partners that are required for signaling. These data are in keeping with observations showing that TLR2 and TLR4, but not IL-1β, signaling is dependent on membrane microdomains, and that after stimulation, TLR2 and TLR4 associate with many proteins within these domains (8, 32, 40, 49, 50, 51, 52, 53). Importantly, the cytokine secretion observed in response to IL-1β revealed that membrane microdomain disruption did not result in complete cellular paralysis, and the lack of toxicity of SAPS with no evidence of impairment of cellular metabolic function suggests that normal uptake mechanisms for environmental sampling and nutrient uptake are unimpaired.

Some oxidized phosphocholine (PC) phospholipids have shown similar properties, with disruption of TLR2 and TLR4 signaling and impairment of TLR4 translocation to lipid rafts/caveolar fractions of endothelial cells (54). However, the inhibitory effects appear to require oxidation of the molecule as the unoxidized form had no effect. It is well documented that oxidized phospholipids induce endothelial cells to interact with monocytes via the production of monocyte-specific chemoattractants (55, 56), adhesion molecules (57), and colony stimulating factors (58). In contrast, SAPS did not require oxidation to function as a potent inhibitor, and we observed only minimal activation of epithelial cells and none of endothelial cells by SAPS, nor did SAPS cause activation of monocyte/ endothelial cells cocultures. Interestingly, however, the inhibitory oxPC phospholipids described by Walton et al. (54) also shared an arachidonoyl group at a similar position on the molecule to SAPS (54). Recent evidence that the nature of the acyl chain structure of membrane lipids is important in the regulation of lipid raft stability (59) suggests that further exploration of the biology of such molecules is warranted. Of note, PS derivatives including SAPS have also been shown to affect secondary Ab production in brown Norway rats previously sensitized to OVA, and although the mechanism remains to be explained, variations of the fatty acid groups altered the subsequent immune response (60).

Additionally, our work revealed a much more profound inhibition of TLR-dependent signaling than just effects on TLR2 and TLR4. While inhibition of individual TLR responses is of interest, it is striking that disruption of membrane microdomains also inhibited responses to heat-killed whole bacteria, which have the potential to engage multiple pattern recognition systems (61). Of particular note, we observed that signaling of a TLR7/8 agonist was markedly impaired in PBMCs and cocultures. This receptor signals solely from acidified endosomes, as does TLR3. In contrast to TLR7/8 signaling, however, SAPS had little impact on TLR3 signaling in epithelial cells. SAPS was active at inhibiting LPS responses in endothelial cell monocultures, and therefore this difference between inhibition of TLR3 and TLR7/8 is unlikely to be a result of global differences between effects of SAPS on leukocytes and tissue cells. The imidazoquinolines such as Gardiquimod are small molecules for which no specific uptake mechanism has been described, and thus it is unlikely that SAPS interfered with Gardiquimod uptake, and global impairment of cellular environmental sampling is unlikely since 1) responses to the much larger, but also intracellular-acting molecule, poly(I:C) were preserved, and 2) the drug had no effect on cellular metabolic capacity. These data reveal that PS species may act as effective membrane disruptors in multiple cellular compartments including the plasma membrane and endosome. Although the role of lipid rafts/membrane microdomains in the signaling of TLRs recognizing viral RNAs has received relatively little attention to date, our results are in keeping with evidence that TLR7/8 shows a greater dependence on membrane microdomains for its signaling than does TLR3 (62, 63). We have not determined the contribution of RIG-I/mda5 (64) to the observed responses to poly(I:C), and if these receptors were still responsive in the face of TLR3 antagonism by SAPS, it is possible that TLR3 signaling has been impaired to a greater extent than our data suggest. Interestingly, oxidized PCs inhibit TLR2 and TLR4 responses but have no effect on responses to TNF-α (54), whose receptor can also reside in lipid rafts (65, 66). To what extent such responses are underpinned by cell-specific signaling systems or interactions of varying phospholipids with specific membrane microdomains showing some potential selectivity for TLRs requires further investigation.

In conclusion, we show herein that effective signaling to a broad range of TLR agonists is mediated by cooperative networks that can be inhibited by a nontoxic phospholipid acting to disturb membrane microdomain formation, and which therefore has the potential to be useful in therapeutic interventions aimed at controlling acute inflammatory responses. The observation that SAPS is one of a variety of forms of PS found in membranes and tissues raises the possibility that its presence in plasma acts as a previously unrecognized autoregulatory antiinflammatory activity.

We thank Miss Kathryn Vaughan and Miss Emily Dick for preparation of the primary human PBMCs, Sue Newton for help with the CBA assay, and Professors Steven K. Dower and Moira K. Whyte for helpful discussions.

The authors have no financial conflicts 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 funded in part by a grant from the Sheffield Hospitals Charitable Trust. I.S. is supported by a MRC Senior Clinical Fellowship (G116/170). K.T. is supported by the Wellcome Trust. J.R.W. is supported by a British Heart Foundation project grant (FS/06/004). This work was also supported by Allergy Therapeutics plc. and Vaccine Technology Ltd. who provided the SAPS (PCT publication no. WO 2008/068621).

3

Abbreviations used in this paper: PS, phosphatidylserine; CBA, cytometric bead array; FRET, fluorescence resonance energy transfer; LTA, lipoteichoic acid; SAPS, 1-stearoyl-2-arachidonoyl-sn-glycero-3-[phospho-l-serine].

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