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. 2017 Jul 13;292(35):14391–14400. doi: 10.1074/jbc.M117.781807

CD14 is a key mediator of both lysophosphatidic acid and lipopolysaccharide induction of foam cell formation

Dong An ‡,§, Feng Hao ‡, Fuqiang Zhang ‡,, Wei Kong §, Jerold Chun ‖, Xuemin Xu ‡, Mei-Zhen Cui ‡,1
PMCID: PMC5582834  PMID: 28705936

Abstract

Macrophage uptake of oxidized low-density lipoprotein (oxLDL) plays an important role in foam cell formation and the pathogenesis of atherosclerosis. We report here that lysophosphatidic acid (LPA) enhances lipopolysaccharide (LPS)-induced oxLDL uptake in macrophages. Our data revealed that both LPA and LPS highly induce the CD14 expression at messenger RNA and protein levels in macrophages. The role of CD14, one component of the LPS receptor cluster, in LPA-induced biological functions has been unknown. We took several steps to examine the role of CD14 in LPA signaling pathways. Knockdown of CD14 expression nearly completely blocked LPA/LPS-induced oxLDL uptake in macrophages, demonstrating for the first time that CD14 is a key mediator responsible for both LPA- and LPS-induced oxLDL uptake/foam cell formation. To determine the molecular mechanism mediating CD14 function, we demonstrated that both LPA and LPS significantly induce the expression of scavenger receptor class A type I (SR-AI), which has been implicated in lipid uptake process, and depletion of CD14 levels blocked LPA/LPS-induced SR-AI expression. We further showed that the SR-AI–specific antibody, which quenches SR-AI function, blocked LPA- and LPS-induced foam cell formation. Thus, SR-AI is the downstream mediator of CD14 in regulating LPA-, LPS-, and LPA/LPS-induced foam cell formation. Taken together, our results provide the first experimental evidence that CD14 is a novel connecting molecule linking both LPA and LPS pathways and is a key mediator responsible for LPA/LPS-induced foam cell formation. The LPA/LPS–CD14–SR-AI nexus might be the new convergent pathway, contributing to the worsening of atherosclerosis.

Keywords: gene regulation, lipid signaling, macrophage, signal transduction, vascular biology, CD14

Introduction

Lysophosphatidic acid (LPA)2 is one of the simplest phospholipids; however, it exerts diverse biological functions on a wide variety of living cells (1–3). The biological effects of LPA are mediated by G protein-coupled receptors: LPA1–3, encoded by endothelial differentiation genes (Edg), and LPA4–6, encoded by a non-Edg family and classified as orphan G protein-coupled receptors belonging to the purinergic receptor family (4–7). LPA can be produced in several ways through the activity of intracellular or extracellular enzymes. The two most prominent pathways involve the conversion of lysophosphatidylcholine to LPA by autotaxin (8, 9) and conversion of phosphatidic acid to LPA by phospholipase A1 or A2 (10, 11). LPA highly accumulates in atherosclerotic lesions (12). Emerging evidence indicates that a long-term, high-fat diet elevates levels of LPA in rabbit and mouse plasma/serum (13, 14). Elevated LPA levels in plasma could affect a divergent function of endothelial cells (ECs), smooth muscle cells (SMCs), monocytes, and macrophages, influencing physiology and pathology of vascular cells and promoting vascular diseases (2).

Macrophages play important roles in all stages of atherosclerosis. The primary origin of macrophages is myeloid progenitor cells in bone marrow (15). Recent lineage-tracing studies reveal that a large portion of macrophage marker-positive cells in mouse and human atherosclerotic lesions are vascular SMC-derived cells (16–18). In early fatty streak lesions, macrophages gather lipoproteins and become lipid-loaded foam cells, playing a crucial role in the development of atherosclerosis. LPA was shown to induce oxidized low-density lipoprotein (oxLDL) uptake in the J774 macrophage cell line (19); however, the underlying molecular mechanism is not well-understood.

Lipopolysaccharide (LPS), usually found in the outer membrane of Gram-negative bacteria, is a potential mediator of inflammatory responses. Chlamydiae-derived chlamydial LPS has been detected in atherosclerotic lesions (20). Repeated intravenous and intraperitoneal administration of LPS accelerates atherosclerosis in rabbits and apoe−/− mice (21–24). The role of common chronic infections in human atherogenesis has also been shown (25). However, the molecular mechanism by which LPS influences atherosclerosis is still not clear. It has been shown that LPS induces macrophage-derived foam cell formation (26, 27). The binding of LPS to its co-receptor, CD14, has been shown to activate inflammatory toll-like receptor pathways (28–30). To date, the role of CD14 in LPS- and LPA-induced oxLDL uptake in macrophages is unknown. In this study, using mouse bone marrow–derived primary macrophages (BMMs), we observed that LPA induces a specific and prominent induction of CD14 mRNA and protein expression and that LPA enhances LPS-induced CD14 expression. Interestingly, LPA also enhances LPS-induced oxLDL uptake in BMMs. We evaluated the role of CD14 in LPA- and LPS-induced oxLDL uptake and discovered a novel role of CD14, which mediates LPA- and LPS-induced oxLDL uptake in BMMs.

This study also determined the molecular mechanism by which CD14 mediates LPA- and LPS-induced oxLDL uptake. Our data revealed that the specific LPA receptor–CD14–scavenger receptor axis mediates LPA- and LPS-induced oxLDL uptake in macrophages. The discovered new mechanism in foam cell formation in macrophages may contribute to atherogenesis.

Results

LPA induces oxLDL uptake and enhances LPS-induced oxLDL uptake in BMMs

Primary mouse macrophages were derived from bone-marrow progenitor cells. The immunofluorescence results indicate that BMMs were positively stained with a CD68 antibody, but not with an α-actin antibody (Fig. 1A). In contrast, SMCs were positively stained with an α-actin antibody but not with a CD68 antibody (Fig. 1A). The purity of BMMs is greater than 95%. Using these BMMs, we observed that LPA time-dependently induced oxLDL uptake (Fig. 1B). Cells were treated with LPA for various times as indicated and then were incubated with Dil-labeled oxidized low-density lipoproteins (Dil-oxLDL) for 3.5 h. Fluorescence data were quantified by a fluorescence plate reader. As shown in Fig. 1B, LPA treatment for 18 h highly induced oxLDL uptake. Our data demonstrated that LPA dose-dependently induced oxLDL uptake in BMMs (Fig. 1C). We observed that LPS also dose-dependently induced oxLDL uptake in BMMs, supporting the same observation in mouse peritoneal macrophages (27). To examine whether LPA affects LPS-induced oxLDL uptake, we stimulated BMMs with various doses of LPA and 100 ng/ml LPS for 18 h, then incubated with Dil-oxLDL for an additional 3.5 h. Interestingly, we found that LPA markedly and dose-dependently enhanced LPS-induced oxLDL uptake in macrophages (Fig. 1C). These results provided the first experimental evidence that LPA enhances LPS-induced oxLDL uptake in macrophage/foam cell formation.

Figure 1.

Figure 1.

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LPA induces oxLDL uptake and enhances LPS-induced oxLDL uptake in BMMs. A, immunofluorescence data revealing the identification of BMMs. BMMs and SMCs were cultured on cover slides overnight. After paraformaldehyde fixation, cells were treated with 0.3% Triton X-100 for permeabilization of the plasma membrane and then immunostained with specific antibodies against CD68, DAPI (nuclear marker), and α-actin. The expression of CD68 (green) and α-actin (red) was examined by Nikon Eclipse E600 fluorescence microscopy; merged images are shown in the right panels. B, LPA-induced oxLDL uptake in BMMs. Starved BMMs were incubated with 5 μm LPA for various time periods (as indicated), followed by the addition of 30 μg/ml DiI-oxLDL for 3.5 h. The uptake of Dil-oxLDL fluorescence value was measured in Synergy HT plate reader at 530 nm excitation and 590 nm emission. The Y axis represents the increased oxLDL uptake levels by LPA compared with the basal oxLDL uptake (uptake of oxLDL alone was considered as 100%). The quantified results were from three independent experiments. *, p < 0.05 versus control. C, the uptake of Dil-oxLDL in BMMs, was dose-dependently induced by LPA and LPS. LPA has an augment effect on LPS-induced Dil-oxLDL uptake. The Y axis represents the increased oxLDL uptake level by LPA, LPS, or LPA plus LPS compared with the basal oxLDL uptake (uptake of oxLDL alone was considered as 100%). Quantified results were from three independent experiments. *, p < 0.05; **, p < 0.01 versus control.

LPA markedly induces CD14 expression in BMMs

To investigate how LPA enhances LPS-induced foam cell formation, we first evaluated whether LPA influences LPS receptor expression in BMMs. CD14 and toll-like receptor 4 (TLR4) are well-characterized co-receptors of LPS (31, 32); TLR2 was also shown to be involved in LPS signaling (33, 34). Therefore, we assessed LPA effect on the expression of these LPS receptors. Cultured mouse BMMs were serum starved for 24 h and then treated with 5 μm LPA for various time periods. Cells were then collected with TRIzol reagent, and the relative CD14 RNA expression levels were evaluated by Northern blotting. As shown in Fig. 2A, we found that LPA significantly induced CD14 transcripts, peaking at around 4 h, but had no effect on either TLR4 or TLR2 RNA expression in macrophages (Fig. 2A). We also observed that LPA dose-dependently induced CD14 protein expression in Western blot analysis (Fig. 2B); peaking at 4 h (Fig. 2C). In the following studies, 5 μm LPA was used because this concentration is in the range of LPA concentrations found in pathological conditions (12). To determine whether LPA-induced CD14 expression was specific for BMMs, we compared the effect of LPA on BMMs with other vascular cell types: SMCs and ECs. As shown in Fig. 2D, LPA-induced CD14 expression in BMMs was specific and significant. LPA does not have a detectable effect on CD14 expression in either SMCs or ECs. As a system control, tissue factor expression in these different cell types was also evaluated (Fig. 2D) because we revealed previously that LPA induced tissue factor expression in SMCs (35). Together, the results presented in Fig. 2 provided the first evidence that LPA markedly and specifically induces the expression of CD14 mRNA and protein in BMMs.

Figure 2.

Figure 2.

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LPA markedly induces CD14 expression in BMMs. A, time course of LPA induction of the RNA expression of LPS receptors. Cultured cells were starved for 24 h prior to 5 μm LPA stimulation for the indicated time periods. Total RNA was extracted using TRIzol reagent and subjected to Northern blot analysis. 18S and 28S RNA were shown as loading controls. B, Western blot analysis showing LPA dose-dependent induction of CD14 expression in BMMs. Cultured cells were starved for 24 h prior to LPA stimulation for various concentrations as indicated. Cell lysates were subjected to Western blot analysis. GAPDH served as the loading control. C, LPA induced CD14 protein expression in BMMs in a time-dependent manner. D, Western blot analysis was performed to evaluate the expression level of CD14 in BMMs compared with vascular SMCs and ECs. The expression of CD14 was detectable and induced by LPA in BMMs. Tissue factor was served as a positive control, which was induced by LPA in SMCs and ECs. GAPDH served as the loading control.

LPA augments LPS-induced CD14 expression in macrophages

It has been shown that LPS up-regulates CD14 expression (36, 37). The effect of LPA on LPS-induced CD14 expression was unknown. We evaluated whether LPA influences LPS-induced CD14 levels. The BMMs were starved for 24 h and then treated with either LPA, LPS, or LPA plus LPS at indicated time points (Fig. 3). Cell lysate was analyzed either with Northern blotting (Fig. 3, A and B) or SDS-PAGE analysis (Fig. 3, C and D). As shown in Fig. 3, stimulation with LPS significantly increased CD14 RNA and protein levels in murine BMMs. Remarkably, we observed that LPA augments LPS effect on CD14 expression at both RNA and protein levels. These data demonstrated for the first time that LPA enhances LPS-induced CD14 expression.

Figure 3.

Figure 3.

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LPA augments LPS-induced CD14 expression in macrophages. A, Northern blot results indicate that LPA enhanced LPS-induced CD14 RNA expression in various time points. Total RNA was extracted using TRIzol reagent and subjected to Northern blot analysis. 18S and 28S RNA were shown as loading controls. The representative Northern blot results shown were from three independent experiments. B, results of the Northern blot analysis were quantified as the densitometry value analyzed by UN-SCAN-IT gel 6.1 software. *, p < 0.05; **, p < 0.01 versus control. C, Western blot results show that LPA increased LPS-induced CD14 protein expression. CD14 protein level was determined by Western blotting with 10% SDS-PAGE. GAPDH served as the loading control. The representative Western blot results shown were from three independent experiments. D, results of the Western blot analysis were quantified as the densitometry value analyzed by UN-SCAN-IT gel 6.1 software. *, p < 0.05; **, p < 0.01 versus control.

CD14 is required for LPA- and LPS-induced oxLDL uptake in BMMs

Results in Fig. 3 show that either LPA or LPS induced CD14 expression, and LPA augmented LPS-induced CD14 expression. Consistent with this pattern, either LPA or LPS induced oxLDL uptake, and LPA augmented LPS-induced oxLDL uptake (Fig. 1). We hypothesized that CD14 might be involved in mediating LPA- and LPS-induced oxLDL uptake in BMMs. To date, it is unknown whether CD14 is involved in LPA or LPS-induced oxLDL uptake/foam cell formation. To explore the role of CD14 in LPA- and LPS-induced oxLDL uptake, we depleted CD14 levels with the specific CD14 siRNA in BMMs and examined the effect of CD14 on oxLDL uptake. Our results demonstrated that the depletion of CD14 largely abolished LPA-, LPS-, or LPA plus LPS–induced Dil-oxLDL uptake in BMMs (Fig. 4A). This conclusion was substantiated by another independent approach: Oil Red O staining (Fig. 4B). Oil Red O staining has been widely used for staining lipids and neutral triglycerides on cells or frozen sections. Together, these data support a new role of CD14 in mediating LPA- and LPS-induced oxLDL uptake; therefore, CD14 is required for LPA- and LPS-induced oxLDL uptake in BMMs.

Figure 4.

Figure 4.

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CD14 is required for LPA- and LPS-induced oxLDL uptake in BMMs. A, knockdown of CD14 expression using the specific CD14 siRNA largely blocked LPA- and LPS-induced oxLDL uptake in BMMs. Non-silencing RNA was used as a negative control. Inset, Western blot results of knockdown of CD14 protein expression. The Y axis represents the increased oxLDL uptake levels by LPA, LPS, or LPA plus LPS compared with the basal oxLDL uptake (uptake of oxLDL alone was considered as 100%). Quantified results were from three independent experiments. *, p < 0.05; **, p < 0.01 versus control; #, p < 0.05; ##, p < 0.01 versus the non-silencing siRNA group. B, effects of LPA, LPS, or LPA plus LPS on oxLDL uptake were examined by the Oil Red O staining with Nikon Eclipse E600 microscopy. BMMs treated with non-silencing RNA or CD14 siRNA (si-CD14) were stimulated with LPA, LPS, or LPA plus LPS for 18 h prior to the treatment of oxLDL for 3.5 h.

Scavenger receptor AI is a downstream mediator of CD14 and is required for LPA- and LPS-induced oxLDL uptake in BMMs

In pursuing the downstream mediator of CD14 for LPA- and LPS-induced oxLDL uptake, we examined the role of scavenger receptor class A type I (SR-AI) because it has been demonstrated that SR-AI was dominantly expressed in mouse macrophages and accounted for 80% of modified LDL uptake (38). However, the regulatory relationship between CD14 and SR-AI in the LPA pathway, the LPS pathway, and an oxLDL uptake has been unknown. We first examined whether LPA or LPS influences SR-AI expression in BMMs. As shown in Fig. 5A, we observed that LPA or LPS increased SR-AI RNA levels time-dependently with the peak around 8 h, and LPA plus LPS has at least an additive effect in BMMs. The quantitative levels of SR-AI are shown in Fig. 5B. The same phenomenon was also observed in protein expression (Fig. 5, C and D). We next examined whether CD14 is required for SR-AI expression in BMMs. Depletion of the CD14 protein with the specific CD14 siRNA largely abolished LPA-, LPS-, and LPA plus LPS–induced SR-AI expression in BMMs (Fig. 5E), indicating that CD14 is the upstream regulator of SR-AI. We then examined the functional role of SR-AI in LPA- and LPS-induced oxLDL uptake. OxLDL uptakes, in response to LPA, LPS, and LPA plus LPS stimulation, were compared between SR-AI antibody-treated and goat IgG-treated BMM groups. We observed that when cell surface SR-AI proteins were quenched by SR-AI–specific antibody, LPA-, LPS-, or LPA plus LPS–induced oxLDL uptakes were completely blocked in comparison to the IgG-treated group (Fig. 5F). Together, these results reveal a new relationship between CD14 and SR-AI in which the CD14–SR-AI pathway mediated LPA-, LPS-, or LPA plus LPS–induced oxLDL uptake/foam cell formation.

Figure 5.

Figure 5.

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SR-AI is a downstream mediator of CD14 and is responsible for LPA- and LPS-induced oxLDL uptake in BMMs. A, RT-PCR results show that LPA and LPS individually induced SR-AI RNA expression and LPA enhanced LPS-induced SR-AI RNA expression at various time points in BMMs. Starved BMMs were stimulated with LPA and LPS for the indicated time periods; the expression levels of SR-AI RNA were determined by RT-PCR for 30 cycles using primer sets specific for SR-AI described in “Experimental Procedures.” Data shown were from three independent experiments. B, RT-PCR results of the SR-AI expression levels were quantified as the densitometry value analyzed by UN-SCAN-IT gel 6.1 software. *, p < 0.05; **, p < 0.01 versus control. C, Western blotting results show that LPA and LPS individually increased SR-AI protein expression; LPA enhanced LPS-induced SR-AI expression in a time-dependent manner. Starved BMMs were stimulated with LPA and LPS for the indicated time periods; cell lysates were examined by Western blot analysis. Equal loading was confirmed by GAPDH loading. Data shown were from three independent experiments. D, results of the Western blot analysis were quantified as the densitometry value analyzed by UN-SCAN-IT gel 6.1 software. *, p < 0.05; **, p < 0.01 versus control. E, knockdown of CD14 protein expression with the specific CD14 siRNA blocked LPA-, LPS-, and LPA plus LPS–induced SR-AI expression. GAPDH served as the loading control. F, effects of the anti-SR-AI antibody on LPA-, LPS-, or LPA plus LPS–induced oxLDL uptake in BMMs. 20 μg/ml SR-AI–specific antibody were used to treat BMMs for 45 min prior to LPA, LPS, or LPA plus LPS treatment. The measurement of oxLDL uptake was described in Fig. 1. Goat IgG was used as a negative control. *, p < 0.05; **, p < 0.01 versus control; #, p < 0.05 versus the Goat-IgG group.

LPA receptor 1 (LPA1) mediates CD14 and SR-AI expression

We next examined which LPA receptor mediates the expression of CD14 and SR-AI. LPA exerts its function on cells through its cognate G protein-coupled receptors. To date, at least six G protein-coupled LPA receptors (LPA1–6) have been reported (39). LPA1–3 share about 50% homology and belong to Edg family G protein-coupled receptors (4–6). A literature search revealed that the LPA Edg family receptors mediate most of the LPA functions in a variety of cell types. To examine the role of Edg family LPA receptors (LPA1, LPA2, and LPA3) in CD14 and SR-AI expression, we first analyzed the expression levels of the LPA Edg family receptors (LPA1–3) in mouse BMMs by RT-PCR. Our data indicated that all LPA1–3 receptors are expressed in BMMs (Fig. 6A). Ki16425, an LPA receptor antagonist with selectivity for LPA1 and LPA3 (40), dose-dependently blocked LPA-induced CD14 expression (Fig. 6, B and C). We then evaluated whether Ki16425 influences LPA plus LPS induction of CD14 expression. As shown in Fig. 6, D and E, 3 μm Ki16425 completely blocked LPA-induced CD14 expression, had a slight effect on LPS-induced CD14 expression, and significantly blocked synergetic induction of LPA and LPS, suggesting a role of LPA1 and LPA3 in CD14 expression. To further identify the specific LPA receptors that mediate LPA function in macrophages, we isolated primary BMMs from wild-type (WT), LPA1, LPA2, and LPA3 knock-out mice (41–44), and examined LPA influence on CD14 expression in these cells. The results showed that only LPA1 deficiency blunted CD14 expression (Fig. 6F), demonstrating that the specific LPA1 is required for LPA-induced CD14 expression. Similarly, LPA1 deficiency prevented LPA-induced SR-AI expression but had no significant effect on LPS-induced SR-AI expression in BMMs (Fig. 6G); LPA1 deficiency diminished the synergetic effect of LPA on LPS-induced SR-AI expression (Fig. 6G). Therefore, LPA1 mediates SR-AI expression. These data support a novel pathway LPA1–CD14–SR-AI axis in live cells.

Figure 6.

Figure 6.

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LPA1 mediates CD14 and SR-AI expression. A, the expression levels of LPA1, LPA2, and LPA3 receptor RNA in BMMs were determined by RT-PCR (30 cycles). Total RNA from BMMs was extracted with TRIzol reagent. After reverse transcription, cDNA was used to perform PCR analysis with mouse LPA1-, LPA2-, and LPA3-specific primers. The RT-PCR results were evaluated in 1.5% agarose gels. DNA markers are indicated on the left side of the gel. B, pretreatment with the LPA1 and LPA3 antagonist Ki16425 dose-dependently blocked LPA-induced CD14 protein expression. Quiescent BMMs were pretreated with Ki16425 at the concentrations indicated for 45 min, and then 5 μm LPA was added for 4 h. CD14 protein levels were determined by Western blotting with 10% SDS-PAGE. GAPDH was used as the loading control. The representative Western blot results shown were from three independent experiments. C, results of the Western blot analysis were quantified as the densitometry value analyzed by UN-SCAN-IT gel 6.1 software. *, p < 0.05 versus control. #, p < 0.05 versus the LPA alone group. D, Ki16425 blocked LPA-induced CD14 expression and the synergistic effect of LPA on LPS induced CD14 expression. GAPDH was used as the loading control. E, results of the Western blot analysis were quantified as the densitometry value analyzed by UN-SCAN-IT gel 6.1 software. *, p < 0.05 versus control; #, p < 0.05 versus LPA, LPS or LPA plus LPS alone group. F, LPA induced CD14 protein expression in WT, LPA1−/−, LPA2−/−, and LPA3−/− BMMs. 5 μm LPA was added to quiescent WT, LPA1−/−, LPA2−/−, and LPA3−/− BMMs for 4 h. CD14 protein level was determined by Western blotting in 10% SDS-PAGE. GAPDH was used as the loading control. G, LPA and LPS induced SR-AI protein expression in WT and LPA1−/− BMMs. LPA1 deficiency abrogated LPA-induced SR-AI protein expression and LPA augment effect on LPS-induced SR-AI protein expression.

LPA1 mediates LPA-induced oxLDL uptake in BMMs

The results in Fig. 6 demonstrate that LPA1 mediates CD14 and SR-AI expression in BMMs. We next examined whether LPA1 mediates LPA induction of oxLDL uptake. Pretreatment with Ki16425 dose-dependently blocked LPA-induced oxLDL uptake (Fig. 7A). Dil-oxLDL fluorescence data were shown, suggesting a role for LPA1 in oxLDL uptake. Using BMMs from either LPA1 knock-out or WT mice, we observed that LPA1 deficiency blocked LPA-induced oxLDL uptake, reduced LPA plus LPS–induced uptake, but had no significant effect on LPS alone–induced uptake (Fig. 7B). Together, the results from Figs. 6 and 7 reveal that the novel LPA1–CD14–SR-AI pathway mediates oxLDL uptake/foam cell formation.

Figure 7.

Figure 7.

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LPA1 mediates LPA-induced oxLDL uptake in BMMs. A, pretreatment with Ki16425, an antagonist for LPA1 and LPA3, dose-dependently blocked LPA-induced oxLDL uptake in BMMs. Ki16425 pretreatment was for 45 min prior to LPA stimulation. The Y axis represents the increased Dil-oxLDL uptake levels by LPA compared with the basal oxLDL uptake (uptake of oxLDL alone was considered as 100%). *, p < 0.05 versus control; #, p < 0.05 versusthe LPA alone group. B, LPA1 deficiency blocked LPA-induced Dil-oxLDL uptake and LPA augment effect on LPS-induced Dil-oxLDL uptake. The Y axis represents the increased oxLDL uptake levels by LPA, LPS, or LPA plus LPS compared with the basal oxLDL uptake (uptake of oxLDL alone was considered as 100%). Quantified results were from three independent experiments. *, p < 0.05 versus control; #, p < 0.05; ##, p < 0.01 versus the WT group.

Discussion

In this study, we made the following novel observations: 1) LPA enhances LPS-induced oxLDL uptake in macrophages; 2) LPA markedly elevates CD14 RNA and protein levels; 3) CD14 is required for LPA- and LPS-induced oxLDL uptake/foam cell formation; 4) CD14 mediates LPA- and LPS-induced SR-AI expression; 5) SR-AI mediates LPA- and LPS-induced foam cell formation; and 6) LPA1 mediates LPA-induced CD14/SR-AI expression and foam cell formation. Taken together, these results demonstrate that CD14 mediates the new convergent pathway of LPA and LPS, leading to biological function in live cells.

LPA accumulates at the atherosclerotic lesions (12, 45). Emerging evidence indicates that a long-term, high-fat diet elevates levels of LPA in animal plasma/serum (13, 14). Elevated LPA levels in plasma could affect divergent functions of ECs, SMCs, and macrophages, influencing physiology and pathology of vascular cells and promoting vascular diseases (2). LPA activates ECs, the innermost layer cells of the vascular wall, by induction of the expression of adhesion molecules (E-selectin, VCAM-1, and ICAM-1) and inflammatory cytokines/chemokines (46, 47). The surface adhesion molecule expression and chemokine secretion by ECs in the vascular wall help recruit monocytes from the bloodstream to the vascular wall to initiate atherosclerotic lesion formation. LPA induces SMC proliferation and migration, which contribute to restenosis and the development of atherosclerosis (48, 49). Our recent studies identified the matricellular protein CCN1 (also called Cyr61), mediating LPA function leading to SMC migration in vitro and in vivo (50, 51). Macrophages play important roles in all stages of atherosclerosis. The primary origin of macrophages is myeloid progenitor cells in bone marrow (15). In early fatty streak lesions, macrophages gather lipoproteins and become lipid-loaded foam cells, which play a crucial role in the development of atherosclerosis. LPA was shown to induce oxLDL uptake in J774 macrophage cell line (19). In this study, we identified a novel pathway, LPA1–CD14–SR-AI, which contributes to macrophage foam cell formation.

The receptors of macrophages help the macrophages sense what is going on, including pathogen-derived molecules (52). CD14 is a glycosylphosphatidylinositol-anchored glycoprotein identified on the surface of monocytes, macrophages, and polymorphonuclear leukocytes (53, 54). CD14 has been shown to be a pivotal membrane receptor for LPS-mediated cellular responses (55, 56). However, whether LPS co-receptor CD14 plays a role in LPA-induced cellular function has been unknown. In this study, we provided the first evidence that LPA via LPA1 significantly induces CD14 expression in macrophages, and that CD14 is a key mediator for both LPA- and LPS-induced oxLDL uptake in macrophages. Therefore, both LPA and LPS pathways converge at CD14 in macrophages for cellular function. Our data also show that CD14 is specifically expressed in macrophages but is not detectable in either mouse aortic ECs or SMCs (Fig. 2). Thus, the LPA-CD14 pathway appears to be specifically important in macrophages.

LPA either enhances or inhibits LPS-induced pathways in various cell types. For instance, LPA enhances LPS-induced IL-6 expression in mouse lung epithelial cell line, but it reduced LPS-induced c-Met tyrosine (Y1003) phosphorylation in human bronchial epithelial cells (57). LPA enhances LPS-induced cyclooxygenase-2 expression in RA synovial cells (58); however, it attenuates LPS-induced cyclooxygenase-2 expression in mouse macrophage cell line J774 (59). These divergent influences of LPA on LPS pathways may depend on LPA-mediated CD14 expression levels in divergent cell types as demonstrated in this study; they may also depend on the interaction of LPA receptors with CD14 on cell membranes as reported previously (60).

TLR4 has been reported as a necessary mediator for LPS-induced oxLDL uptake, promoting foam cell formation (27). Although it was well-documented that CD14 interacts with LPS and other LPS receptor complex components, such as TLR4 and MD-2 to transduce LPS signal, the role of CD14 in LPS-induced oxLDL uptake/foam cell formation was not revealed previously (61). Our results demonstrated that CD14 is an inducible key mediator for LPS-induced foam cell formation. Significantly our data also reveal that CD14 is a key mediator required for LPA-induced oxLDL uptake/foam cell formation. Therefore, CD14 is an essential and convergent mediator, controlling both LPA and LPS pathways.

Scavenger receptor class A (SR-A) has been shown to contribute to the uptake of modified LDL (38, 62) and the development of atherosclerotic lesions (63). Extensive oxidation of LDL appears to be required for rapid uptake via SR-A, whereas mildly oxidized LDL is preferentially internalized via CD36 (64, 65). The identified LPA1-mediated SR-AI induction via CD14 presents a new pathway, which regulates SR-AI function.

Recent evidence has shown a role for gut microbiota in atherosclerosis. Besides the ability of bacterially derived metabolites to act as hormones modulating cardiovascular risk, gut hyperpermeability (leaky gut) allows bacterial cell wall products such as LPS to enter into the bloodstream to activate macrophages and modulate risk of developing atherosclerosis (66, 67). Both LPA and LPS have been found in atherosclerotic lesions (12, 20). In this study, our data show that LPA synergizes LPS effect.

Taken together, our data provide the first experimental evidence that CD14 connects both LPA and LPS pathways, leading to biological function-macrophage foam cell formation. As macrophage uptake of oxLDL plays an important role in foam cell formation and the pathogenesis of atherosclerosis, the LPA/LPS–CD14–SR-AI nexus identified in this study might be a new convergent pathway contributing to the worsening of atherosclerosis.

Experimental procedures

Reagents

Lysophosphatidic acid (LPA) was purchased from Avanti Polar Lipids (Alabaster, AL). Lipopolysaccharide (LPS) was obtained from Sigma-Aldrich. TRIzol reagent was from Invitrogen. Antibodies against mouse CD14, SR-AI, and tissue factor were from R&D Systems (Minneapolis, MN). Antibody against GAPDH was from EMD Millipore (Billerica, MA). Antibody against CD68 was from BioLegend (San Diego, CA). Antibody against smooth muscle α-actin was from Thermo Fisher Scientific. GoTaq Flexi DNA Polymerase and the reverse transcription system were from Promega (Madison, WI). The RNeasy kit, non-silencing siRNA, and CD14 siRNA were from Qiagen (Gaithersburg, MD). Primers for LPA receptors and oxLDL receptor SR-AI used for conventional PCR were as follows: LPA1, 5′-AGC TGC CTC TAC TTC CAG C-3′ (forward) and 5′-TTG CTG TGA ACT CCA GCC AG-3′ (reverse); LPA2, 5′-ATG GGC CAG TGC TAC AAC G-3′ (forward) and 5′-AGG GTG GAG TCC ATC AGT G-3′ (reverse); LPA3, 5′-GAC AAG CGC ATG GAC TTT-3′ (forward) and 5′-CAT GTC CTC GTC CTT GTA CG-3′ (reverse); SR-AI, 5′-AAA ATG GCC CCT CCG TTC AG-3′ (forward) and 5′-ATC CGC CTA CAC TCC CCT TCT C-3′ (reverse). Human LDL and Dil-labeled oxLDL were purchased from Biomedical Technologies Inc. (Stoughton, MA).

Cell culture

Bone marrow progenitor cells were harvested from the femur section of 8- to 10-week-old C57B/6 mice, which were from The Jackson Laboratory (Bar Harbor, ME). After 6 consecutive days of culture in DMEM (20% M-CSF and 10% fetal bovine serum) more than 95% of bone marrow progenitor cells were differentiated into macrophages. The M-CSF–conditioned medium was prepared by collecting the supernatant from 10% serum DMEM cultured LADMAC cells (ATCC, Manassas, VA) and filtering through a 0.22 μm filter (Millipore).

RT-PCR analysis

RNA expression levels of various LPA receptors and SR-AI were evaluated. Total RNA was isolated from BMMs using TRIzol reagent. The first strand of cDNA was reverse transcribed. The cDNA products were amplified using GoTaq Flexi DNA Polymerase. Amplification conditions were as follows: 5 min at 95 °C and 30 cycles of 30 s at 95 °C, 30 s at 55 °C, and 1 min at 72 °C. The reaction was followed by a final extension for 10 min at 72 °C. The PCR products were analyzed by electrophoresis on a 1.5% agarose gel.

Western blot analysis

Cultured mouse BMMs were rinsed with cold PBS and lysed in Western blot lysis buffer (50 mm Tris-HCl, pH 6.8, 8 m urea, 5% mercaptoethanol, 2% SDS, and protease/phosphatase inhibitors) with sonication for 30 s on ice. Cellular proteins were separated by 10% SDS-PAGE and transferred to a polyvinylidene fluoride membrane (Immobilon-P, Millipore). Membranes were then probed with the specific antibodies, and the specific protein bands were viewed using ECL Plus (GE Healthcare).

Northern blot analysis

Total cellular RNA was isolated using TRIzol reagent according to the manufacturer's instructions. Total RNA (8–10 μg) was subjected to denaturing electrophoresis on formaldehyde-agarose gels. RNA was blotted onto Amersham Hybond nylon membrane (GE Healthcare) and hybridized with 32P-labeled cDNA probes. 18S and 28S ribosomal RNA were used as internal controls.

siRNA treatment

BMMs were transfected with non-silencing or specific siRNA (Qiagen) for 48 h, using Lipofectamine RNAiMAX Reagent (Thermo Fisher Scientific) following the instructions provided by the manufacturer. On day 3, cells were cultured in serum-free medium for 24 h, followed by treatment either with or without inducers.

Oil Red O staining

BMMs were cultured on microscope cover glasses in 12-well plates and starved for 24 h. After the LPA and LPS treatment, the cells were incubated with oxidized LDL at 37 °C for 3.5 h. LDL was oxidized by dialysis against 10 μm CuSO4 in PBS for 24 h at room temperature. After the completion of treatments, cells were rinsed once with 1× PBS, fixed in 4% paraformaldehyde at 4 °C for 30 min. Subsequently, the cells were rinsed once with 1× PBS and then stained with Oil Red O solution (Sigma Aldrich) at room temperature for 10 min. After staining, the cells were rinsed two times with 1× PBS then stained with hematoxylin (Sigma Aldrich) for 1 min. Afterward, they were rinsed two times with 1× PBS and observed by light microscopy (Nikon Eclipse E600 microscope).

Analysis of Dil-oxLDL uptake

BMMs were cultured in 24-well plates and starved for 24 h. After an LPA and LPS treatment for 18 h, cells were incubated with Dil-labeled oxLDL (Biomedical Technologies) for 3.5 h and then rinsed with cold PBS containing 5% BSA two times. After further rinsing with PBS for two more times, the cells were detached from the culture plate with 5% Triton X-100 in PBS. The fluorescence value was measured by the Synergy HT plate reader (BioTek, Winooski, VT) at excitation 530 nm and emission 590 nm.

Immunofluorescence

BMMs grown on slide cover glass were fixed in 4% ice cold paraformaldehyde solution for 30 min followed by treatment with 0.3% Triton X-100 in PBS for 5 min at room temperature. The cells were then incubated for 1 h in 5% goat serum blocking buffer (Sigma) plus 0.1% Tween 20 in PBS and incubated with CD68 antibody or smooth muscle α-actin in 1/200 dilution overnight at 4 °C. After being washed with PBS three times (5 min each), the cells were incubated with the secondary antibody, goat anti-sheep IgG Alexa Fluor 488, or Rhodamine Red-X-AffiniPure Goat Anti-Mouse IgG for 2 h at room temperature. Then the cells were washed with PBS four times (5 min each) at room temperature, incubated with DAPI for 2 min, and washed with PBS three times (5 min each) at room temperature. Subsequently, the cover glasses were mounted on slides with permanent aqueous mounting medium (BioGenex, Fremont, CA), and the labeled cells were analyzed by fluorescence microscopy with a Nikon Eclipse E600 microscope.

Statistical analysis

Results are means ± S.E. Comparisons between multiple groups were performed using one-way analysis of variance with post hoc t tests. Single comparisons were made using two-tailed, unpaired Student's t tests. A p value of 0.05 was considered statistically significant.

Author contributions

M.-Z. C. conceived the idea and coordinated the research. D. A., F. H., and F. Z. designed and performed experiments and analyzed data. D. A. prepared the figures. J. C. provided LPA receptor knock-out mice. D. A., X. X., K. W., J. C., and M.-Z. C. analyzed and interpreted data. D. A. and M.-Z. C. wrote the manuscript. All authors edited, revised, and approved the final version of the manuscript.

This work was supported by National Institutes of Health Grants HL107466 (to M.-Z. C.) and NS095256 (to X. X.). This work was also supported by the University of Tennessee Center of Excellence in Livestock Diseases and Human Health. The authors declare that they have no conflicts of interest with the contents of this article. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

2
The abbreviations used are:
LPA
lysophosphatidic acid
oxLDL
oxidized low-density lipoprotein
BMM
bone marrow–derived macrophage
M-CSF
macrophage colony–stimulating factors
SMC
smooth muscle cell
EC
endothelial cell
TLR2
toll-like receptor 2
TLR4
toll-like receptor 4
SR-AI
scavenger receptors class A type I
LPA1
LPA receptor 1.

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