Critical role of peroxisome proliferator-activated receptor α in promoting platelet hyperreactivity and thrombosis under hyperlipidemia.

Platelet hyperreactivity and increased atherothrombotic risk are specifically associated with dyslipidemia. Peroxisome proliferator-activated receptor alpha (PPARα) is an important regulator of lipid metabolism. It has been suggested to affect both thrombosis and hemostasis, yet the underlying mechanisms are not well understood. In this study, the role and mechanism of PPARα in platelet activation and thrombosis related to dyslipidemia were examined. Employing mice with deletion of PPARα (Pparα-/-), we demonstrated that PPARa is required for platelet activation and thrombus formation. The effect of PPARα is critically dependent on platelet dense granule secretion, and is contributed by p38MAPK/Akt, fatty acid b-oxidation, and NAD(P)H oxidase pathways. Importantly, PPARα and the associated pathways mediated a prothrombotic state induced by a high-fat diet and platelet hyperactivity provoked by oxidized low density lipoproteins. Platelet reactivity was positively correlated with the levels of expression of PPARα, as revealed by data from wild-type, chimeric (Pparα+/-), and Pparα-/- mice. This positive correlation was recapitulated in platelets from hyperlipidemic patients. In a lipid-treated megakaryocytic cell line, the lipid-induced reactive oxygen species-NF-kB pathway was revealed to upregulate platelet PPARα in hyperlipidemia. These data suggest that platelet PPARα critically mediates platelet activation and contributes to the prothrombotic status under hyperlipidemia.

Introduction

Underpinned by platelet hyperactivity, atherothrombotic disease is the leading cause of mortality and morbidity worldwide. Dyslipidemia has been firmly established as a risk factor for atherothrombotic disease.[1,2] Despite the vigorous efforts that have been devoted to establishing the pathways leading to platelet hyperactivity in dyslipidemia,[3,4] the mechanisms responsible are still unclear. Identification of key targets by which dyslipidemia regulates platelet activity is imperative for the prevention and management of atherothrombotic disease.

Oxidized low density lipoproteins (oxLDL), the product of dysfunctional lipid metabolism, are major promoters of a prothrombotic state in both animal models and human patients.[5,6] Scavenger receptor CD36 and signaling pathways such as Src family kinases (SFK), mitogen-activated protein kinases and reactive oxygen species (ROS) are involved in oxLDL-induced platelet activation.7- [9] Molecules involved in lipid metabolism such as the transcription factors farnesoid X receptor,[10] liver X receptor,[11] and PPAR[12-14] are also expressed in platelets. How these molecules interact with the established platelet activation network is ill-defined.

PPARα is a major regulator of lipid metabolism in nucleated cells by upregulating the transcription of lipid-metabolizing enzymes, such as carnitine palmitoyl-CoA transferase-I (CPT-I) and acyl-CoA oxidase.[15-17] It is expressed in anucleate platelets and was reported to play roles in thrombosis and hemostasis.[18,19] Although PPARα may interact with protein kinase C during platelet activation,[18] the underlying signaling mechanism has not been elucidated. A variety of metabolic and pathological conditions are related to PPARα expression,[20-22] but it has not been explored whether or how platelet activation is regulated by PPARα.

In this study, we investigated the role of PPARα in dyslipidemia-related prothrombotic potential and platelet hyperactivity. PPARα expression in platelets was enhanced in both a hyperlipidemic mouse model and patients, which correlated well with platelet hyperactivity. The mechanism increasing PPARα expression in platelets and the platelet functions targeted by PPARα were explored.

Methods

Subjects

The procedures in human subjects were approved by the Ethics Committee of the First Affiliated Hospital of Zhejiang University, and informed consent was obtained from the study participants. Blood from 36 patients with hypertriglyceridemia, 16 patients with hypercho lesterolemia and 31 healthy subjects was obtained. None of the participants had taken any antiplatelet or other nonsteroidal anti-inflammatory drugs for at least 14 days before blood collection.

None of the patients had clinical evidence of cardiovascular disease (according to their clinical history, physical examination, and electrocardiogram). Moreover, exclusion criteria for all subjects included renal insufficiency, proteinuria, altered hepatic function and alcohol abuse. Patients with diabetes mellitus (fasting blood glucose level >115 mg/dL or treatment with a hypoglycemic agent), hypertension (systolic blood pressure >140 mmHg, diastolic blood pressure >90 mmHg) and smokers were also excluded.

Animals

Previously reported PPARα-deficient mice (Ppara) were used in these experiments.[23] All mice were 8 to 14 weeks old, and matched for weight and sex. Male ApoE-deficient (Apoe) mice (6 weeks old) were purchased from the Model Animal Research Center of Nanjing University (Nanjing, China). Ppara mice were generated by crossing Ppara and Apoe mice in the animal facilities of Zhejiang University. The animals were fed a normal chow diet until 8 weeks. Their diet was then switched to a high-fat diet, containing 40% fat and 1.25% cholesterol (Trophic Animal Feed High-Tech Co., Ltd, China), for 8 weeks. All animals were maintained under standard conditions of room temperature and humidity with a 12-hour dark-light cycle. All animal protocols were approved by Zhejiang University Laboratory Animal Welfare and Ethics Committee.

Preparation of washed human platelets

All blood donors had antecubital veins allowing a clean venipuncture. Blood was drawn without stasis into siliconized vacutainers containing 1/9 v/v 3.8% sodium citrate, then washed platelets were re-suspended as previously described.[24]

Preparation of washed mice platelets

Whole blood was collected from the inferior vena cava into a 0.2 volume of ACD buffer (75 mM sodium citrate, 39 mM citric acid, and 135 mM dextrose, pH 6.5), and was diluted 1:3 with modified Tyrode buffer (20 mM HEPES, 137 mM NaCl, 13.8 mM NaHCO3, 2.5 mM KCl, 0.36 mM NaH2PO4, 5.5 mM glucose, pH 7.4). Diluted whole blood was centrifuged at 180 g for 10 min at room temperature. The platelet-rich plasma was collected into a fresh tube containing 500 mL ACD, and centrifuged at 700 g for 10 min. The platelet pellet was then re-suspended in modified Tyrode buffer.

Statistical analysis

Results are expressed as mean ± standard error of the mean (SEM). Statistical significance was evaluated with a paired t-test, two-tailed Mann-Whitney U tests and two-way analysis of variance (ANOVA) using the statistical software GraphPad Prism (GraphPad Software, La Jolla, CA, USA).

Results

Pparα mice display impaired hemostasis and thrombosis

Ppara mice were genotyped by polymerase chain reaction and ablation of Ppara was confirmed in PPARα-deficient platelets by western blot analysis (Figure 1A). The levels of expression of PPARβ and PPARγ were similar in heterozygous (Ppara) platelets, Ppara platelets and wild-type (WT) platelets (Figure 1A). Moreover, Pparamice were viable and fertile, and did not exhibit any evident bleeding tendency or thrombotic events over their lifespan. Ppara mice did not differ significantly from their WT littermates with regard to platelet count, red blood cell count, white blood cell count, hematocrit, or hemoglobin concentration (Online Supplementary Table S1). Electron microscopy showed normal discoid morphology of Ppara platelets with unaltered numbers of a granules and dense granules, compared to those in the WT platelets (Online Supplementary Figure S1A). No significant differences in the surface expression of platelet CD41 (aIIb subunit), and CD42b (GPIbα subunit) were found between WT and Ppara platelets (Online Supplementary Figure S1B).

Figure 1.

PPARα-deficient mice display impaired hemostasis and thrombosis. (A) Genotyping results of wild-type (WT), Pparaand Ppara mice using polymerase chain reaction. Immunoblot analysis of PPARα, PPARβ and PPARγ expression in platelets from WT, Ppara mice and humans. (B) Bleeding times for WT (•), Ppara (■) and Ppara (▲) mice. Means are indicated by horizontal lines. Statistical significance was evaluated with a paired t test (*P<0.05; ***P<0.001; ns: not significant). Percentages of WT, Ppara and Ppara mice bleeding times that exceeded 15 min (D) or were within 15 min (■). Results were obtained from 26 WT, 26 Ppara and 26 Ppara mice. (C) An injury to the carotid artery was induced by FeCl3. The dot plot shows occlusion times for carotid arterioles as a result of FeCl3-induced thrombosis in WT (•, n=6), Ppara (■, n=9) and Ppara mice (▲, n=6). Means are indicated by horizontal lines. Statistical significance was evaluated with a two-tailed Mann-Whitney test (*P<0.05; **P<0.01; ns: not significant). (D) Representative images and time courses of thrombus formation induced by FeCl3 injury to mesenteric arterioles in WT (top row), Ppara (middle row) and Ppara (bottom row) mice. a: arteriole; v: venule. Scale bars, 100 mm (left panel). Dot plot showing occlusion times for arterioles as a result of FeCl3-induced thrombosis in WT (•, n=22), Ppara (■, n=33) and Ppara mice (▲, n=26). Means are indicated by horizontal lines. Statistical significance was evaluated with a two-tailed Mann-Whitney test (*P<0.05; ns: not significant). (E) Photomicrographs showing the progression of adhesion of platelets from WT, Ppara and Ppara mice on collagen. Whole blood from WT, Ppara and Ppara mice, collected in heparin (7.5 U/mL), was fluorescently labeled by incubation with mepacrine (100 mM) for 30 min, and then perfused through fibrillar collagen-coated bioflux plates at a shear rate of 40 dynes/cm[2] for 5 min. Original magnification, ×10. Scale bar, 100 mm (left panel). Dot plot showing area coverage of platelets from WT (•), Ppara (■) and Ppara (▲) mice (n=3 for each group; two-way analysis of variance test, *P<0.05; ***P<0.001).

With a tail-bleeding assay, Ppara mice showed significantly prolonged tail bleeding time (712.80 ± 58.11 seconds vs. 333.40 ± 64.76 seconds; P<0.001) (Figure 1B), agreeing with findings from a previous study.[18] Moreover, 73% of the Ppara mice had a bleeding time exceeding 15 min, while the percentage in WT littermates was 19% (Figure 1B). In a FeCl3-induced model of carotid artery thrombosis, the time to formation of a stable occlusive thrombus in the carotid artery was significantly longer in Ppara mice than in WT mice (10.88 ± 1.51 min vs. 3.53 ± 0.59 min, P<0.01) (Figure 1C). In a FeCl3-induced model of mesenteric arteriole thrombosis, the time to formation of stable occlusive thrombi was significantly longer in Ppara mice than in WT mice (38.35 ± 3.10 min vs. 27.55 ± 2.81 min, P<0.05) (Figure 1D). Interestingly, heterozygous Ppara+/-mice also had a significant perturbance of thrombotic and hemostatic functions, with significantly increased tail bleeding time, rate of non-stoppable bleeding, and time to the formation of stable occlusive thrombi compared to those of WT mice (Figure 1B-D). In a model of deep vein thrombosis, Ppara+/- and Ppara mice developed thrombi similar in weight and length to those observed in WT mice (Online Supplementary Figure S2A, B), indicating a complex multi-cellular interaction upon PPARα deficiency in thrombo-inflammation. These in vivo data indicate that PPARα is essential for hemostasis and thrombosis, functions governed by platelets; nevertheless, the role of PPARα is complicated in thrombo-inflammation because the outcome is dictated by the interaction of platelets and inflammatory cells.

In a microfluidic perfusion assay, when whole-blood was perfused over an immobilized collagen surface at the shear stress of 1000 s-1 for 5 min, the areas covered by Ppara+/- and Ppara platelets were 27.5% and 44.9% smaller in average than those by WT platelets (Figure 1E). A recombinant whole-blood system with diluted washed platelets (2 × 107/mL) was used in the same assay to assess the collagen-adhesion ability of platelets. In the absence of platelet aggregation, the areas covered by collagen-adhered Ppara platelets were similar to those of WT platelets (Online Supplementary Figure S3). These findings indicate that PPARα functions in regulating the growth of platelet thrombi, not the initial adhesion.

Pparα platelets show functional defects due to an impaired ATP secretion

Next, platelet aggregation in response to common platelet stimuli was analyzed. Compared to WT platelets, Pparaplatelets displayed an average 30% reduction of aggregation rates in response to thrombin (0.025 U/mL) and 57% reduction in response to collagen (0.8 mg/mL) (Figure 2A). However, ADP and TXA2 analog U46619-induced platelet aggregation was not affected by PPARα deficiency (Online Supplementary Figure S4A). Although dense granule content was normal in Ppara platelets (Online Supplementary Figure S4B), ATP release induced by low doses of thrombin (0.025 U/mL), collagen (0.8 mg/mL) and U46619 (0.3 mM) was largely inhibited in Ppara platelets (Figure 2A and Online Supplementary Figure S4A). Again, Ppara+/-platelets exhibited intermediate rates of aggregation and dense granule secretion (Figure 2A). Higher concentrations of thrombin (0.05 U/mL) and collagen (2 mg/mL) overcame the defective aggregation and dense granule secretion in Ppara platelets (Online Supplementary Figure S4A). The aggregation differences between WT and Ppara platelets were abolished when apyrase was applied to hydrolyze dense granule-secreted ATP and ADP (Figure 2B and Online Supplementary Figure S4C). Conversely, supplementation with a low concentration of ADP (1 mM), which was insufficient to induce aggregation on its own, rescued the defective aggregation of Ppara+/- and Ppara platelets stimulated by thrombin or collagen (Figure 2C and Online Supplementary Figure S4D). As indicated by the measurement of TXB2, collagen- or thrombin-induced TXA2 production was comparable between WT and Pparaplatelets (Online Supplementary Figure S4E). Moreover, the secretion of a-granules and activation of aIIbb3, measured respectively by P-selectin expression and the binding of Jon/A antibody, were not influenced by PPARα deficiency in response to thrombin and convulxin (Online Supplementary Figure S4F). These data suggest that the impaired aggregation in Ppara platelets is caused by the reduced ADP secretion. Consistent with the role of ADP in thrombus amplification, Ppara platelets formed smaller aggregates than WT platelets when stimulated with low doses of thrombin and collagen (Online Supplementary Figure S4G).

Figure 2.

Ppara (A) Aggregation and ATP release of washed wild-type (WT), Ppara and Ppara platelets stimulated with thrombin (0.025 U/mL) or collagen (0.8 mg/mL). Aggregation and ATP release were assessed with a Chrono-log lumiaggregometer under stirring at 1,200 rpm. Traces are representative of at least three independent experiments. Statistical significance was evaluated with a two-tailed Mann-Whitney test and a paired t test (*P<0.05; **P<0.01; ***P<0.001). (B) Aggregation and ATP release of washed WT, Ppara and Pparaplatelets stimulated with thrombin (0.025 U/mL) or collagen (0.8 mg/mL) in the presence of vehicle or apyrase (1 U/mL) incubated for 5 min. Traces are representative of at least three independent experiments. (C) Aggregation of washed WT, Ppara and Pparaplatelets stimulated with thrombin (0.025 U/mL) or (0.8 mg/mL) in the presence of a low concentration of ADP (1 mM). Traces are representative of at least three independent experiments. (D) Spreading of WT, Ppara and Pparaplatelets on immobilized fibrinogen in the presence or absence of apyrase (1 U/mL) or ADP (1 mM). Images are representative of three independent experiments with similar results. Original magnification, ×100. Scale bar, 10 mm (left panel). Statistical significance was evaluated with a two-tailed Mann-Whitney test (**P<0.01; ***P<0.001; ns: not significant). (E) Platelets from WT, Pparaand Pparamice were resuspended with human platelet-poor plasma at a concentration of 4×108/mL, and recombined plasma was stimulated to coagulate with thrombin (0.4 U/mL), then photographed at different time points. Statistical significance was evaluated with a paired t test (*P<0.05; **P<0.01).

Platelet spreading on immobilized fibrinogen and clot retraction, two processes controlled by early and late integrin aIIbb3-mediated outside-in signaling, respectively, were then measured. Platelet spreading on immobilized fibrinogen (Figure 2D) and clot retraction (Figure 2E) were also inhibited by PPARα deficiency. Apyrase eliminated the spreading difference between WT and Ppara platelets, and exogenous ADP (1 mM) rescued the defective spreading of Ppara platelets (Figure 2D). Clot retraction mediated by Ppara platelets showed a significant delay compared to that by WT platelets (Figure 2E). These data demonstrate an important role for PPARα in platelet dense granule secretion and its activation.

PPARα promotes platelet activation through a p38/ROS/Akt signal axis

When platelet signaling events were analyzed, both collagen and thrombin induced a significantly reduced phosphorylation of Akt (Thr308/Ser473) and p38 (Thr180/Tyr182) in Ppara platelets (Figure 3A and Online Supplementary Figure S5A), while phosphorylation of ERK1/2 (Thr202/Tyr204) and JNK (Thr183/Tyr185) remained unaltered (Online Supplementary Figure S5A). Moreover, both SB203580 and SH-6, inhibitors of p38 and Akt, strongly inhibited platelet aggregation and ATP release induced by thrombin and collagen, but no additive effects with PPARα deficiency were observed (Figure 3B and Online Supplementary Figure S5B). Therefore, PPARα is functionally coupled to p38 and Akt activation. As p38 regulates the production of ROS,[25,26] thrombin and convulxin induced significantly less ROS production in Ppara+/- and Ppara platelets than in WT platelets (Figure 3C), while ROS scavenging by N-acetylcysteine (NAC) essentially eliminated the aggregation and ATP release difference among WT, Ppara+/-, and Pparaplatelets (Figure 3D and Online Supplementary Figure S5C).

Figure 3.

PPARα promotes platelet activation through a p38/ROS/Akt signal axis. (A) Immunoblot analysis of wild-type (WT), Pparaand Pparaplatelets, stimulated with thrombin (0.025 U/mL) and collagen (0.8 mg/mL) for 5 min, with antibodies recognizing phosphorylated Akt Thr308, phosphorylated Akt Ser[47] [3], total Akt, phosphorylated p38 Thr180/Tyr182 (T180/Y182), and total p38. Representative immunoblots from at least three independent experiments. (B) Washed WT, Ppara and Pparaplatelets (2×108/mL) were incubated with dimethyl sulfoxide (DMSO), SH-6 (10 mM), SB203580 (10 mM) for 10 min, then stimulated with thrombin (0.025 U/mL) or collagen (0.8 mg/mL), respectively. Aggregation and ATP release were assessed with a Chrono-log lumiaggregometer under stirring at 1,200 rpm. Traces are representative of at least three independent experiments. (C) Generation of reactive oxygen species (ROS) analyzed by flow cytometry. H2DCFDA-loaded (50 mM) mice platelets were stimulated with thrombin (0.025 and 0.05 U/mL) or convulxin (50 and 100 ng/mL) for 5 min. Samples were analyzed immediately. Statistical significance was evaluated with a two-tailed Mann-Whitney test (*P<0.05; **P<0.01; ns: not significant). (D) Washed WT, Ppara and Pparaplatelets (2×108/mL) were incubated with or without N-acetylcysteine (NAC, 2 mM) for 5 min, then stimulated with thrombin (0.025 U/mL) or collagen (0.8 mg/mL), respectively. Aggregation and ATP release were assessed with a Chrono-log lumiaggregometer under stirring at 1,200 rpm. Traces are representative of at least three independent experiments. (E) Immunoblot analysis of WT, Ppara and Pparaplatelets stimulated with thrombin (0.025 U/mL) and collagen (0.8 mg/mL) for 5 min in the absence or presence of NAC, with antibodies recognizing phosphorylated Akt Thr308, phosphorylated Akt Ser[47] [3], total Akt, phosphorylated p38 Thr180/Tyr182 (T180/Y182), and total p38. Representative immunoblots from at least three independent experiments. (F) Immunoblot analysis of WT, Ppara and Pparaplatelets, stimulated with thrombin (0.025 U/mL) or collagen (0.8 mg/mL) for 5 min in the presence of DMSO, SH-6 (10 mM), and SB203580 (10 mM), with antibodies recognizing phosphorylated Akt Thr308, phosphorylated Akt Ser[47] [3], total Akt, phosphorylated p38 Thr180/Tyr182 (T180/Y182), and total p38. Representative immunoblots from at least three independent experiments. (G) H2DCFDA-loaded (50 mM) mice platelets were incubated with DMSO, SH-6 (10 mM) or SB203580 (10 mM), stimulated with thrombin (0.05 U/mL) or convulxin (100 ng/mL) for 5 min. Samples were analyzed immediately. Statistical significance was evaluated with a two-tailed Mann-Whitney test (*P<0.05; **P<0.01; ***P<0.001; ns: not significant). (H) Washed WT, Ppara and Pparaplatelets (2×108/mL) were incubated with DMSO, etomoxir (25 mM), VAS2870 (10 mM), or ALP (200 mM) for 10 min, then stimulated with thrombin (0.025 U/mL) or collagen (0.8 mg/mL), respectively. Aggregation and ATP release were assessed with a Chrono-log lumiaggregometer under stirring at 1,200 rpm. Traces are representative of at least three independent experiments.

Intriguingly, NAC treatment abolished the difference of Akt phosphorylation but left intact the difference of p38 phosphorylation among WT, Ppara+/- and Ppara (Figure 3E and Online Supplementary Figure S6A). Furthermore, SB203580 and SH-6 eliminated the differences of Akt phosphorylation (Figure 3F and Online Supplementary Figure S6B) and ROS production (Figure 3G) between WT and Ppara platelets; whereas SH-6 did not change the phosphorylation of p38 (Figure 3F and Online Supplementary Figure S6B). These data suggest a sequential relay of PPARα, p38, ROS production, and Akt during platelet activation.

Sources of ROS were also investigated using the NADPH oxidase inhibitor VAS2780, CPT-I inhibitor etomoxir, and xanthine oxidase inhibitor allopurinol (Online Supplementary Figure S6C). VAS2780 and etomoxir, but not allopurinol, eliminated the differences of platelet aggregation and ATP release between WT and Ppara platelets (Figure 3H and Online Supplementary Figure S6D). Both VAS2780 and etomoxir eliminated the phosphorylation difference of Akt, but not that of p38 between WT and Ppara platelets (Online Supplementary Figure S6E). These data suggest that ROS from NADPH oxidase and mitochondrial fatty acid b-oxidation constitute the important sources of ROS for PPARα-regulated platelet activation.

Consistent with previous reports,[18,19] a synthetic PPARα agonist WY14643 inhibited aggregation and ATP release induced by low doses of thrombin and collagen in WT platelets (Online Supplementary Figure S7A), and essentially abolished the differences between WT and Ppara platelets (Online Supplementary Figure S7A). Unexpectedly, a PPARα antagonist GW6471 also inhibited aggregation and ATP release in WT platelets and eliminated the difference between WT and Ppara platelets in response to low doses of thrombin and collagen (Online Supplementary Figure S7B). WY14643 and GW6471 both inhibited phosphorylation of Akt Ser[47] [3] upon platelet activation by collagen, indicating that these compounds act by interrupting PPARα signaling in platelets (Online Supplementary Figure S7C). Moreover, GW6471 per se but not WY14643 induced phosphorylation of Akt Ser[47] [3] (Online Supplementary Figure S7D). Therefore, GW6471 may be a partial agonist in the context of platelet activation. Hence, the agonist and antagonist of PPARα both appear to inhibit platelet function, possibly due to the disruption of signal transduction mediated by PPARα.

PPARα mediates hyperlipidemia-associated prothrombotic status and oxidized low-density lipoprotein-evoked platelet activation

Consistent with the previous study,[27] after 8 weeks of a high-fat diet, total plasma levels of cholesterol and triglycerides in Ppara/Apoe-/- mice were significantly increased, while they did not undergo further change in Ppara/Apoe mice (Online Supplementary Table S2). The occlusion time in FeCl3-induced mesenteric arteriole thrombosis in Ppara/Apoe mice was significantly shortened by a high-fat diet when compared with the control diet. But it was comparable between Ppara/Apoe mice fed with a high-fat diet and Ppara/Apoe mice fed the control diet (Figure 4A).

Figure 4.

PPARα mediates hyperlipidemia-associated prothrombotic status and oxLDL-evoked platelet activation. (A) Ppara/Apoe-/- and Ppara/Apoe-/- mice were fed a high-fat diet (HFD) or control diet (CD) for 8 weeks before undergoing in vivo thrombosis experiments. Platelets were labeled by direct tail vein injection of DiOC6 (10 mM, 100 mL). Dot plot showing occlusion times for arterioles as a result of FeCl3-induced thrombosis in Pparaand Ppara/Apoe mice. Means are indicated by horizontal lines. Statistical significance was evaluated with a two-tailed Mann-Whitney test (*P<0.05; **P<0.01). (B) Platelets were stimulated with 1-(palmitoyl)-2-(5-keto-6-octenedioyl) phosphatidylcholine (KODiA-PC, 15 mM). Aggregation was assessed with a Chrono-log lumiaggregometer under stirring at 1,200 rpm. Traces are representative of at least three independent experiments. (C) Immunoblot analysis of wild-type (WT), Ppara and Pparaplatelets stimulated with KODiA-PC for 5 min, with antibodies recognizing phosphorylated Src Tyr418 (Y418), total Src, phosphorylated p40phox, b-actin, phosphorylated ERK5 Thr218/Tyr220 (T218/Y220), total ERK5, phosphorylated p38, total p38, phosphorylated Akt Ser473, and total Akt (top panel). Statistical significance was evaluated with a paired Student t test (*P<0.05; **P<0.01; ***P<0.001) (bottom panel). (D) Washed WT, Ppara and Pparaplatelets were incubated with N-acetylcysteine (NAC, 2 mM), etomoxir (Eto, 25 mM), dimethylsulfoxide (DMSO), VAS2870 (10 mM), BIX02188 (10 mM), SB203580 (10 mM), or SH-6 (10 mM) for 10 mins, then stimulated with KODiA-PC (15 mM). Aggregation was assessed with a Chrono-log lumiaggregometer under stirring at 1,200 rpm. Traces are representative of at least three independent experiments. (E) H2DCFDA-loaded (50 mM) mice platelets were incubated with DMSO, NAC (2 mM), VAS2870 (10 mM), or etomoxir (25 mM), stimulated with KODiA-PC (15 mM) for 10 min. PAPC was used as a negative control. Samples were analyzed immediately. Statistical significance was evaluated with a two-tailed Mann-Whitney test (*P<0.05; ns: not significant).

KODiA-PC, 1-(palmitoyl)-2-(5-keto-6-octene-dioyl) phosphatidylcholine, one of the most potent CD36 ligands in the oxLDL species, caused direct platelet aggregation, which was largely decreased in Ppara platelets (Figure 4B and Online Supplementary Figure S8A). Consistently, KODiA-PC induced a significantly reduced phosphorylation of Src (Tyr418), p40phox and ERK5 (Thr218/Tyr220), Akt (Ser473) and p38 (Thr180/Tyr182) in Ppara platelets, compared with WT platelets (Figure 4C). Moreover, the NADPH oxidase inhibitor VAS2780, the ROS scavenger NAC, the ERK5 inhibitor BIX02188, the p38 inhibitor SB203580 and the Akt inhibitor SH-6, but not the CPT-I inhibitor etomoxir, eliminated the difference in aggregation between WT and Ppara platelets (Figure 4D and Online Supplementary Figure S8B). Robust ROS production induced by KODiA-PC was also reduced by PPARα deficiency (Figure 4E). Interestingly, ROS production by KODiA-PC was inhibited by VAS2780 and NAC, but not etomoxir (Figure 4E). These data indicate that PPARα mediates oxLDL-induced platelet activation, which is associated with an altered ROS generation pathway.

Platelet PPARα expression correlates with platelet hyperreactivity

Having established the importance of PPARα in hyperlipidemic-induced platelet activation, we further found that PPARα expression in platelets was significantly increased in mice fed a high-fat diet, compared to that in mice fed with a control diet (Figure 5A). The expression of platelet PPARβ and PPARγ was however not significantly influenced by the high-fat diet (Figure 5A). In mice fed a high-fat diet, the increase of PPARα expression was accompanied by an increase in platelet aggregation induced by thrombin and collagen (Figure 5B and Online Supplementary Figure S9). These data indicate that increased platelet PPARα expression in hyperlipidemic mice is responsible for platelet hyperactivity.

Figure 5.

Increased platelet PPARα expression correlates with platelet hyperreactivity in hyperlipidemic mice and patients with hyperlipidemia. (A) Immunoblot analysis of PPARα, PPARβ and PPARγ expression in platelets from Apoe mice fed a high-fat diet (HFD) or control diet (CD) for 8 weeks with PPARα, PPARβ and PPARγ antibodies. Statistical significance was evaluated with a two-tailed Mann-Whitney test (**P<0.01; ns: not significant). (B) Aggregation and ATP release of platelets from Apoe mice fed with a HFD or CD were stimulated with thrombin (0.015 U/mL) or collagen (0.6 mg/mL). Aggregation was assessed with a Chrono-log lumiaggregometer under stirring at 1,200 rpm. Traces are representative of at least three independent experiments. (C) Immunoblot analysis of PPARα, PPARβ and PPARγ expression in platelets from patients with hypertriglyceridemia (HTG) and hypercholesterolemia (HTC) with PPARα, PPARβ and PPARγ antibodies. Representative immunoblots of platelet PPARα and glyceraldehyde 3-phosphate dehydrogenase (GAPDH) from four healthy subjects, six patients with HTG and three patients with HTC. Representative immunoblots of platelet PPARβ, PPARγ and GAPDH from four healthy subjects, three patients with HTG and three patients with HTC. Statistical significance was evaluated with a two-tailed Mann-Whitney test (*P<0.05; ns: no significance). (D) PPARA mRNA expression in platelets from healthy subjects (n=10) and patients with HTG (n=12) or healthy subjects (n=9) and patients with HTC (n=10) was analyzed by quantitative real-time polymerase chain reaction. Statistical significance was evaluated with a two-tailed Mann-Whitney test (*P<0.05). (E) Aggregation of platelets from healthy subjects (n=25) and patients with HTG (n=36) or healthy subjects (n=16) and patients with HTC (n=16) in response to thrombin (0.025 U/mL). Aggregation was assessed with a Chrono-log lumiaggregometer under stirring at 1,200 rpm. Statistical significance was evaluated with a two-tailed Mann-Whitney test (*P<0.05). (F) Platelet aggregation induced by thrombin is well correlated with protein level of platelet PPARα expression in healthy subjects (n=25) and patients with HTG (n=36) or healthy subjects (n=16) and patients with HTC (n=16). Each solid circle represents a different individual (Pearson correlation, GraphPad Prism 5).

Platelet PPARα expression was also determined in healthy volunteers and patients with hyperlipidemia (Online Supplementary Table S3). Platelet PPARα protein and mRNA levels were significantly increased in patients with hypertriglyceridemia and hypercholesterolemia (Figure 5C, D), although the expression of PPARβ and PPARγ proteins was similar in healthy subjects and hyperlipidemic patients (Figure 5C). As expected, platelet aggregation in response to thrombin was enhanced in patients with hypertriglyceri-demia or hypercholestero lemia compared to that in healthy subjects (Figure 5E). The level of platelet PPARα expression was closely correlated with platelet aggregation in response to thrombin (Figure 5F). These findings demonstrate that the increased expression of platelet PPARα in patients with hyperlipidemia is closely related to platelet activity.

Oxidized low-density lipoproteins and lipids upregulate megakaryocyte- but not platelet- PPARα

Compared with platelets treated with normal medium, platelets incubated with a fatty acid (oleic acid or palmitic acid), cholesterol or oxLDL for 12 h or 24 h did not alter the PPARα levels (Online Supplementary Figure S10A). In contrast, PPARα protein and mRNA levels in Meg-01 cells were significantly increased after 24 h incubation with the fatty acid, cholesterol or oxLDL (Figure 6A, B and Online Supplementary Figure S10B), without a concomitant change of the expression of PPARβ and PPARγ (Figure 6A and Online Supplementary Figure S10B). These data suggest that the increased PPARα in hyperlipidemic platelets may derive from megakaryocytes.

Figure 6.

Oxidized low-density lipoproteins and lipids upregulate megakaryocyte but not platelet PPARα. (A) Immunoblot analysis of PPARα, PPARβ and PPARγ expression in Meg-01 cells cultured with fatty acids (oleic acid [OA], 400 mM and palmitic acid [PA], 200 mM), cholesterol (CHO, 2.5 mg/mL, 5.0 mg/mL, 7.5 mg/mL) or oxidized low-density lipoproteins (oxLDL, 10 mg/mL, 50 mg/mL) for 24 h with PPARα, PPARβ and PPARγ antibodies. Representative immunoblots from at least three independent experiments. (B) PPARA mRNA expression in Meg01 cells cultured with fatty acids (OA, 400 mM and PA, 200 mM), cholesterol (CHO, 2.5 mg/mL, 5.0 mg/mL, 7.5 mg/mL) or oxLDL (10 mg/mL, 50 mg/mL) for 24 h was analyzed by quantitative real-time polymerase chain reaction. Statistical significance was evaluated with a two-tailed Mann-Whitney test (*P<0.05; ns: not significant). (C) Immunoblot analysis of PPARα and phosphorylated IκBα level in Meg-01 cells cultured with fatty acids (OA, 400 mM and PA, 200 mM), cholesterol (CHO, 2.5 mg/mL, 5.0 mg/mL, 7.5 mg/mL) or oxLDL (10 mg/mL, 50 mg/mL) in the absence or presence of BAY11-7082 (10 mM) for 24 h. Representative immunoblots from at least three independent experiments. (D) Immunoblot analysis of PPARα and phosphorylated IκBa level in Meg-01 cells cultured with fatty acids (OA, 400 mM and PA, 200 mM), CHO (2.5 mg/mL, 5.0 mg/mL, 7.5 mg/mL) or oxLDL (10 mg/mL, 50 mg/mL) in the absence or presence of N-acetylcysteine (NAC, 1 mM) or DTT (1 mM) for 24 h. Representative immunoblots from at least three independent experiments. (E) NF-kB binding to the Ppara promoter of Meg-01 cells as determined by chromatin immunoprecipitation. Schematic diagram showing the NF-kB-binding site in the Ppara promoter (top panel). Amplification of the Ppara promoter region containing the NF-kB-binding motif in Meg-01 cells cultured with fatty acids (OA, 400 mM and PA, 200 mM), CHO (2.5 mg/mL, 5.0 mg/mL, 7.5 mg/mL) or oxLDL (10 mg/mL, 50 mg/mL) for 24 h. GAPDH was used as a control to show precipitation specificity (bottom panel). Results shown are representative of three or more separate experiments run on different days.

It was reported that hyperlipidemia induces ROS generation and activation of the NF-kB signaling pathway.[28] Indeed, treating Meg-01 cells with the NF-kB inhibitor BAY11-7082, antioxidants NAC or DTT, abolished the PPARα upregulation by fatty acids, cholesterol or oxLDL (Figure 6C, D and Online Supplementary Figure S10C). Moreover, BAY11-7082 and NAC or DTT inhibited fatty acid-, cholesterol- or oxLDL-induced phosphorylation of IκBα (Figure 6C, D and Online Supplementary Figure S10C). An in silico promoter analysis (Jaspar and ensemble Genome Brower) identified the possible NF-kB-binding sites on the Ppara promoter. Four potential NF-kB sites in the sense strand of the region -105/-114 bp (region 1), -168/-177 bp (region 2), -1588/-1597 bp (region 3) and -1878/-1887 bp (region 4). Chromatin immunoprecipitation analyses revealed a marked binding of p65 to region 1 of the Ppara promoter of megakaryocytes when treated with fatty acid, cholesterol or oxLDL (Figure 6E), indicating that NF-kB directly regulates Ppara transcription in Meg-01 cells. Thus, fatty acids, cholesterol or oxLDL upregulate PPARα expression in Meg01 cells through ROS and subsequent NF-kB signaling.

Discussion

The present study investigated the role of PPARα in platelet activation and the impact of PPARα in the prothrombotic potential caused by hyperlipidemia. The results demonstrated that PPARα is an indispensable signaling molecule supporting platelet activation and thrombosis. Importantly, increased PPARα expression in platelets is responsible for enhanced platelet activity by hyperlipidemia. Hyperlipidemia does not trigger PPARα expression in platelets directly, but rather does so in megakaryocytes through ROS and NF-kB pathways. Our study not only elucidated the signaling function of PPARα in supporting platelet activation, but also revealed a key role for PPARα in bridging the genetic effect of hyperlipidemia on megakaryocytes with the prothrombotic potential operated by platelets.

This study clearly demonstrated the positive role that PPARα serves in supporting platelet activation and thrombosis. However, previous studies showed some synthetic[18,19] or endogenous molecules[19] inhibited platelet activation and thrombosis in a PPARα-dependent manner. This discrepancy may suggest the existence of endogenous PPARα ligands which serve as a positive regulator of platelet function and thrombosis. Although such ligands of PPARα have yet to be defined, given the phenotype of platelets upon PPARα deficiency, it is possible that these stimulatory ligands are the ones playing dominant roles in platelet activation. While lipids with cardioprotective effects (e.g., polyunsaturated fatty acids)[29] are able to produce endogenous PPARα ligand with platelet-inhibitory properties (e.g., DPAn-6),[19] lipid species with cardiovascular disease-promoting properties, such as saturated fatty acids,[29] may generate derivatives which serve as stimulatory PPARα ligands to promote platelet activation. Identification of the stimulatory ligands may thus provide novel targets for the intervention of thrombosis and constitutes an important theme in its own right. Moreover, based on the fact that ligands mainly target the ligand-binding domain of PPARα,[30,31] it is tempting to hypothesize that a conformational change induced by occupancy of the region of the ligand-binding domain may be key to the non-genomic function of PPARα. Future structural studies are therefore imperative to provide further insight into this promising anti-thrombotic target.

This study not only confirmed that the key contribution of PPARα to thrombosis and hemostasis is through the regulation of platelet dense granule secretion, but also revealed the pathway on which PPARα relies to perform its function. Hence, p38 and Akt were identified as the sequential signals downstream of PPARα to regulate platelet dense granule secretion. These findings are in agreement with the previously reported roles of p38 and Akt in platelets. For example, p38 has been shown to positively regulate dense granule secretion,[25] and Akt have also been reported to be important in dense granule secretion through promoting nitric oxide/cGMP signaling[32] or inhibiting GSK3β.[33] Moreover, our results suggest that p38 activation is relayed by Akt phosphorylation, which is also consistent with a previous report.[34] Given the intact Jon/A binding and the significantly reduced spreading on immobilized fibrinogen upon PPARα deficiency, it seems that PPARα mainly participates in outside-in signaling rather than inside-out signaling. This observation is also consistent with previous reports, which suggested that outside-in signaling and dense granule secretion are coupled and both are regulated by p38 and Akt signaling, evidenced by the studies employing either inhibition or genetic ablation of p38 or Akt.[25,32] Downstream of p38, ROS generation has been found to be a pivotal link between PPARα and platelet activation, echoing a previous finding in macrophages.[35] It seems that hemostatic stimuli[36-38] and oxLDL[9] may differentially employ the ROS generation pathways. Notably, our data indicate that PPARα critically contributes to the generation of ROS controlled by both mitochondrial fatty acid β-oxidation and NADPH oxidases. Therefore, the PPARα/p38/ROS/Akt axis may function as a central gatekeeper for platelet activation and is employed by major platelet receptors, such as immunoreceptor tyrosine-based activation motif receptor, G-protein-coupled receptors, and possibly CD36.

A key finding of the present study is the correlation between the level of expression of PPARα and the extent of platelet reactivity. The subsequent data from the hyperlipidemic Pparα/Apoe mice model further revealed a causative relationship between PPARα and platelet hyperactivity under the condition of hyperlipidemia. The correlation is found in both humans and mice, which indicates that it might be an evolutionary conservative mechanism. Unlike in macrophages, where oxLDL elevates the expression of both PPARα[21] and PPARγ,[39] megakaryocytes seems to respond to hyperlipidemia specifically with upregulated expression of PPARα, but not of PPARβ or PPARγ. Considering the importance of fatty acids in supporting both megakaryocyte maturation[40] and platelet production,[41] and the central roles of PPARα in fatty acid metabolism,[17,42] it may not be surprising that megakaryocyte PPARα is a sensitive and specific target regulated by hyperlipidemia. Although it is commonly accepted that nongenomic mechanisms direct PPAR family-mediated platelet function,[13,18,43,44] our findings suggest that in the case of PPARα, a genomic regulation in megakaryocytes and a nongenomic signaling role in platelets are concerted to underscore the platelet hyperreactivity under hyperlipidemia.

In the current study, NF-kB, activated by ROS, was revealed as the possible mechanism of the upregulation of PPARα in megakaryocytes by hyperlipidemia. Our data suggest that NF-kB upregulates PPARα expression possibly through direct binding to the Ppara promoter region in lipid-treated megakaryocytes. It is worth mentioning that hyperglycemia elicits a similar NF-κB activation which subsequently upregulates the expression of P2Y12, a key receptor in mediating platelet activation and thrombosis.[45] It is therefore possible that metabolic disorders activate common inflammatory pathways to promote thrombosis. Interestingly, it was reported that PPARα acts as a feedback to negatively regulate NF-kB activation in smooth muscle cells[46] and the metabolism of the arachidonic acid metabolite leukotriene B4 in hepatocytes.[47] It is yet to be determined whether PPARα acts similarly in megakaryocytes to negatively regulate inflammation. However, while upregulation of PPARα may suppress inflammation, it may exert a prothrombotic effect through enhanced platelet activation. PPARα may thus be a key molecule maintaining the balance of thrombosis and inflammation, as suggested by the thrombo-inflammatory model of deep vein thrombosis in the current study.

The study has several limitations. First, expression of PPARα in vascular cells has been reported;[48-50] in the absence of a cell-specific knockout model, a possible contribution to hemostasis from vasculature PPARα could not be excluded. Second, functional validation of the signaling involved was performed with inhibitors, which inevitably have possible off-target effects. Although it is beyond the scope of the study to chase every off-target effect, care must be taken when interpreting the results.

To summarize, we found that platelet PPARα positively mediates platelet activation through promoting dense granule secretion. Hyperlipidemia may contribute to the prothrombotic status through upregulated expression of PPARα in megakaryocytes/platelets and enhanced activation signaling mediated by PPARα in platelets. This work suggested that coupled genomic and nongenomic interventions targeting PPARα may be necessary for the prevention of thrombosis under hyperlipidemia.