Alzheimer's disease is associated with the accumulation of Aβ (amyloid β)-peptides in the brain. Besides their cytotoxic effect on neurons, Aβ-peptides are thought to be responsible for the atherothrombotic complications associated with Alzheimer's disease, which are collectively known as cerebrovascular disease. In the present study, we investigated the effect of Aβ-peptides on human platelet signal transduction and function. We discovered that the 25–35 domain of Aβ-peptides induce an increase in platelet intracellular Ca2+ that stimulates α-granule and dense granule secretion and leads to the release of the secondary agonist ADP. Released ADP acts in an autocrine manner as a stimulant for critical signalling pathways leading to the activation of platelets. This includes the activation of the protein kinases Syk, protein kinase C, Akt and mitogen-activated protein kinases. Ca2+-dependent release of ADP is also the main component of the activation of the small GTPase Rap1b and the fibrinogen receptor integrin αIIbβ3, which leads to increased platelet aggregation and increased thrombus formation in human whole blood. Our discoveries complement existing understanding of cerebrovascular dementia and suggest that Aβ-peptides can induce vascular complications of Alzheimer's disease by stimulating platelets in an intracellular Ca2+-dependent manner. Despite a marginal ADP-independent component suggested by low levels of signalling activity in the presence of apyrase or P2Y receptor inhibitors, Ca2+-dependent release of ADP by Aβ-peptides clearly plays a critical role in platelet activation. Targeting ADP signalling may therefore represent an important strategy to manage the cerebrovascular component of Alzheimer's disease.

INTRODUCTION

Abnormal metabolism of the APP (amyloid precursor protein) through the amyloidogenic pathway results in the accumulation of heterogeneous Aβ (amyloid β)-peptides (40–42 amino acids) in the central nervous system, causing the onset of AD (Alzheimer's disease) [1,2]. Amyloid peptides accumulate also in the cerebrovascular system of AD patients and play a role in the onset and progression of cerebrovascular diseases, such as stroke and CAA (cerebral amyloid angiopathy) [3]. Aβ-peptides are responsible for the prothrombotic state described in several AD patients, where increased clot formation, decreased fibrinolysis and elevated levels of coagulation factors are responsible for reduced blood flow to the brain, neuroinflammation and ultimately neurodegeneration [4]. Platelets are the major source of amyloid peptides found in the plasma. Platelets express APP and the enzymatic machinery responsible for its metabolism and for the generation of Aβ-peptides [57]. Under physiological conditions platelets metabolize APP through the non-amyloidogenic pathway: α-secretase is activated by Ca2+ and calmodulin [8] and releases sAPPα (soluble APPα) that regulates thrombosis and haemostasis in vivo [9]. In AD, however, β- and γ-secretase activity increases, and APP is metabolized to produce Aβ-peptides, which leads to alteration of the APP content of platelets [10]. sAPP fragments and Aβ-peptides are stored in platelet α-granules and released upon stimulation with physiological agonists [11]. Several studies have suggested that platelet-derived Aβ-peptides may represent a reliable peripheral biomarker of AD [12,13]. Abnormalities in platelet membrane fluidity and secretase activity, as well as increased APP level, have been documented in platelets from AD patients [14].

It has become increasingly clear that Aβ-peptides directly activate platelets. Herczenik et al. [15] showed that misfolded amyloid peptides induce platelet activation and aggregation through two different pathways initiated by CD36 or GPIb (glycoprotein Ib) respectively. A short peptide corresponding to the biologically active sequence of Aβ (Aβ25–35) [16,17] is able to activate human platelets and potentiate platelet aggregation induced by low doses of collagen, ADP and thrombin through a PI3K (phosphoinositide 3-kinase) and p38MAPK (mitogen-activated protein kinase)-dependent pathway [18,19]. Sonkar et al. [20] have recently demonstrated that Aβ25–35 stimulates platelet activation through RhoA-dependent modulation of actomyosin organization. Moreover, immobilized Aβ-peptides are able to induce platelet adhesion and activation of the intracellular signalling pathway, and to fasten spreading over collagen [21]. Despite these reports, the exact mechanism responsible for Aβ-peptide-dependent activation of platelets remains elusive.

In the present study, we investigated further the molecular mechanism of Aβ-induced platelet activation. One of the most interesting hypotheses on the mechanism for Aβ toxicity is based on the ability of the peptides to form cation-permeable pores in the cellular membranes, which ultimately results in membrane depolarization, Ca2+ leakage, and disruption of ionic homoeostasis [22,23]. Because of the effect of Aβ-peptides on intracellular Ca2+ in other cell types [24], we investigated the effect of this protein on platelet intracellular Ca2+ homoeostasis. We demonstrated that Aβ-peptides stimulate robust Ca2+ mobilization and granule secretion. Despite some evidence for marginal levels of ADP-independent platelet activation, ADP release resulting from dense granule secretion plays a critical role in stimulating canonical signalling pathways, which include PI3K, PKC (protein kinase C), p38MAPK and ERK (extracellular-signal-regulated kinase), as well as tyrosine kinases, which include Syk kinase. These events support platelet aggregation and thrombus formation in human whole blood under physiological shear stress. Taken together, our data unequivocally identify Ca2+-dependent release of ADP as the molecular mechanism underlying the pro-thrombotic effect of Aβ-peptides and further clarifies the link between AD and cerebrovascular disease.

EXPERIMENTAL

Materials

25–35, BSA, DMSO, prostaglandin E1, indomethacin, apyrase grade I and VII, collagen type I, leupeptin, aprotinin, RGDS (Arg-Gly-Asp-Ser), MRS-2179, BAPTA [1,2-bis-(o-aminophenoxy)ethane-N,N,N′,N′-tetra-acetic acid] and calcein/AM (acetoxymethyl ester) (green) were purchased from Sigma–Aldrich. AR-C69931MX was a gift from AstraZeneca. SB202580, PD98059, Y27632 and Ly294002 were from Alexis. Ro31-8220 was from Calbiochem. The rabbit polyclonal antibodies against Rap1 (121) and Syk (N19) and the monoclonal antibody anti-tubulin (DM1A) were from Santa Cruz Biotechnology. Anti-[phospho-Akt (Ser473)], anti-[phospho-p38MAPK (Thr180/Tyr182)], anti-[phospho-ERK1/2 (Thr202/Tyr204)], anti-[phospho-MLC (myosin light chain) (Ser19)] and anti-(PKC phospho-substrates) antibodies were from Cell Signaling Technology. Appropriate peroxidase-conjugated anti-IgG antibodies were from Bio-Rad Laboratories. The ECL substrate and fura 2/AM were from Millipore. FITC-labelled fibrinogen was from Molecular Probes. FITC-conjugated human P-selectin was from Beckman Coulter. The ATP determination kit was from Biaffin.

Preparation of washed human platelets

Human platelets were obtained from healthy volunteers, in accordance with the Declaration of Helsinki of the World Medical Association and approved by the Ethics Committee of the University of Pavia, as described previously with minor modifications [25]. Briefly, whole blood in citric acid/citrate/dextrose (152 mM sodium citrate, 130 mM citric acid and 112 mM glucose) was centrifuged at 120 g for 10 min at room temperature. Apyrase grade I (0.2 units/ml), prostaglandin E1 (1 μM) and indomethacin (10 μM) were then added to PRP (platelet-rich plasma). Platelets were recovered by centrifugation at 720 g for 15 min, washed with 10 ml of Pipes buffer (20 mM Pipes and 136 mM NaCl, pH 6.5), and finally gently resuspended in Hepes buffer (10 mM Hepes, 137 mM NaCl, 2.9 mM KCl and 12 mM NaHCO3, pH 7.4). The cell count was typically adjusted to 5×108 platelets/ml unless otherwise stated.

Analysis of platelet activation

Unless otherwise stated, platelet samples were typically stimulated with 10–20 μM Aβ25–35 or 0.1 units/ml thrombin as a control. Aggregation of platelets (2×108 platelets/ml, 0.25 ml) was monitored continuously in a Born lumiaggregometer for up to 5 min. Rap1b activation was analysed by a pull-down assay using the GST-tagged Rap-binding domain of RalGDS (GST–RalGDSRBD) essentially as described previously, and precipitated active Rap1b was identified by immunoblotting with an anti-Rap1 antibody [26]. Measurement of cytosolic Ca2+ was performed on 0.4 ml samples of fura 2/AM-loaded platelets under gentle stirring in a PerkinElmer Life Sciences LS3 spectrofluorimeter in the presence of 1 mM CaCl2 or 2 mM EGTA, according to a previously published procedure [27]. For analysis of protein phosphorylation, identical amounts of proteins from whole platelet lysates were separated by SDS/PAGE on 5–15% or 10–20% acrylamide gradient gels, and transferred on to PVDF. Immunoblotting analysis was performed as described previously [28], using the following antibodies and dilutions: anti-Syk (N-19); anti-[phospho-Akt (Ser473)], anti-[phospho-p38MAPK (Thr180/Tyr182)], anti-[phospho-ERK1/2 (Thr202/Tyr204)] and anti-[phospho-MLC (Ser19)], all at 1:500 dilution; anti-(PKC phospho-substrates), anti-tubulin (DM1A) and anti-Rap1 (121), all at 1:1000 dilution. Reactive proteins were visualized by ECL. All of the immunoblots are representative of at least three different experiments giving similar results.

Flow cytometry

Samples of washed platelets (106 cells in 0.05 ml of Hepes buffer containing 1 mM CaCl2, 1 mM MgCl2 and 0.2% BSA), untreated or activated with different doses of Aβ25–35 were labelled for 10 min at room temperature with different specific antibodies: FITC-conjugated anti-P-selectin or FITC-conjugated fibrinogen. The reaction was stopped by diluting samples with 0.45 ml of Hepes buffer. Samples were immediately analysed by flow cytometry using a FACSCalibur instrument equipped with CellQuest Pro software (BD Bioscience). Data analyses were performed using FlowJo 7.6.1 software (Tree Star).

Measurement of [14C]serotonin secretion and ATP/ADP release

Agonist-induced release of [14C]serotonin from metabolically labelled platelets was performed as described previously [27]. Platelet secretion was determined by measuring the release of ATP by adding luciferin-luciferase reagent. Briefly, 5×108 platelets/ml (0.3 ml) were stimulated with the indicated concentrations of Aβ25–35, and reactions were stopped by adding 10 mM EDTA. Cells were spun down and 50 μl of supernatant were analysed with the ATP determination kit (Biaffin) following the manufacturer's instructions.

Thrombus formation assay

The Bioflux200 system (Fluxion) was used to analyse thrombus formation in human whole blood under flow essentially as described previously [29,30]. Microchannels were coated for 30 min with 1 mg/ml collagen I (monomeric collagen from calf skin), before blocking with 0.5% BSA and washing with modified Tyrode's buffer. The PRP was isolated from blood by centrifugation (200 g, 15 min) and incubated for 1 h with Calcein™ (5 μg/ml) at 37°C to facilitate thrombus visualization in whole blood. The PRP was then added to the red blood cell fraction to reconstitute whole blood with physiological blood cell density (including platelets), and thrombus formation (on collagen) was visualized by fluorescence microscopy. Where indicated, 20 μM Aβ25–35 and/or 2 units/ml apyrase was added to the PRP 10 min before the start of the flow assay. Platelet adhesion under flow conditions was tested on collagen I (shear rate=1000 s−1). Representative pictures were taken at time 10 min and surface area coverage for the entire microchannel was measured using Bioflux200 Software (version 2.4).

Statistical analysis

Data are expressed throughout as means±S.E.M. The results were analysed by Student's t test (for comparisons of two groups) or one-way ANOVA with Bonferroni post-test (for multiple comparisons).

RESULTS

25–35 induces intracellular Ca2+ increase and granule release

Several studies demonstrate that the toxicity of Aβ-peptides resides in their ability to increase intracellular Ca2+ by forming cation-permeable pores [22]. The ability to form ionic pores has been demonstrated for amyloid peptides of different length and is associated with the Aβ25–35 domain [31]. Accordingly, the Aβ25–35 domain has been shown to be responsible for the neurotoxic properties of Aβ-peptides [16] and has been widely used to investigate the biological properties of Aβ-peptides in different cell types, including platelets [1821]. In the present study, we analysed the direct effect of Aβ25–35 on Ca2+ mobilization in washed human platelets labelled with fura 2/AM, both in the presence and in the absence of extracellular Ca2+. Representative traces reported in Figure 1(A) show platelet Ca2+ movements in the presence of 1 mM extracellular CaCl2 upon stimulation with thrombin or Aβ25–35. Aβ25–35 (20 μM) induced a slow increase in Ca2+ with a kinetic profile different from the spike observed in thrombin-stimulated platelets. Pre-treatment of platelets with EGTA, which chelates extracellular Ca2+, completely abolished Ca2+ movements. Quantitative analysis shown in Figure 1(B) reveals that Aβ25–35 promotes a statistically significant increase in intracellular Ca2+. Recent evidence suggested a pivotal role for RhoA in platelet activation induced by Aβ25–35 and the activity of ROCK (Rho-associated kinase) was found necessary for Aβ25–35-induced cytoskeletal rearrangements and aggregation [20]. Therefore we tested the effect of the ROCK inhibitor Y27632 on the Aβ25–35-induced increase in intracellular Ca2+ and demonstrated that Ca2+ influx is upstream of RhoA activation in the signalling event stimulated by Aβ25–35 since the inhibition of this kinase did not significantly affect Ca2+ influx (Figure 1C). In addition, despite a non-statistically significant trend towards reduction in Ca2+ response in the presence of the ADP-degrading enzyme apyrase, the inhibition of ADP signalling does not significantly inhibit intracellular Ca2+ movements induced by Aβ25–35 (Figure 1D). This suggests that the effect of Aβ25–35 on intracellular Ca2+ is mostly ADP-independent.

25–35 induces inward Ca2+ in washed platelets

Figure 1
25–35 induces inward Ca2+ in washed platelets

Platelets (2×108/ml, 0.4 ml) were loaded with fura 2/AM (3 μM, 40 min) and stimulated with 1 unit/ml thrombin or 20 μM Aβ25–35 in the presence of 1 mM extracellular CaCl2 or 2 mM EGTA. (A) Traces of Ca2+ mobilization are representative of three different experiments. (B) Histograms show the quantification of the intracellular concentration of Ca2+ 4 min after stimulation with 20 μM Aβ25–35 (closed bars, 1 mM extracellular CaCl2; open bar, 2 mM EGTA). (C) Platelets were stimulated with 20 μM Aβ25–35 and where indicated pre-incubated with 20 μM Y27632. (D) Platelets were stimulated with 20 μM Aβ25–35 and where indicated pre-incubated with 2 units/ml apyrase for 10 min. Histograms show the quantification of the intracellular concentration of Ca2+ 4 min after stimulation. Statistical analysis was performed using one-way ANOVA with Bonferroni post-test, n=3, *P<0.05 and **P<0.01. ns, P>0.05.

Figure 1
25–35 induces inward Ca2+ in washed platelets

Platelets (2×108/ml, 0.4 ml) were loaded with fura 2/AM (3 μM, 40 min) and stimulated with 1 unit/ml thrombin or 20 μM Aβ25–35 in the presence of 1 mM extracellular CaCl2 or 2 mM EGTA. (A) Traces of Ca2+ mobilization are representative of three different experiments. (B) Histograms show the quantification of the intracellular concentration of Ca2+ 4 min after stimulation with 20 μM Aβ25–35 (closed bars, 1 mM extracellular CaCl2; open bar, 2 mM EGTA). (C) Platelets were stimulated with 20 μM Aβ25–35 and where indicated pre-incubated with 20 μM Y27632. (D) Platelets were stimulated with 20 μM Aβ25–35 and where indicated pre-incubated with 2 units/ml apyrase for 10 min. Histograms show the quantification of the intracellular concentration of Ca2+ 4 min after stimulation. Statistical analysis was performed using one-way ANOVA with Bonferroni post-test, n=3, *P<0.05 and **P<0.01. ns, P>0.05.

We next sought to determine platelet granule secretion. In the presence of 1 mM CaCl2, Aβ25–35 induced a dose-dependent α-granule release, measured as P-selectin exposure (Figure 2A) and dense granule release, measured as [14C]serotonin secretion (Figure 2B). Platelet dense granule release was also evaluated by measuring the extracellular accumulation of ADP/ATP using the luciferin/luciferase assay. We found that platelets at the physiological density of 5×108/ml released ADP at concentrations of 0.146±0.1 and 0.184±0.13 μM in response to 10 μM and 20 μM Aβ25–35 respectively. As shown in Figure 2(B), dense granule secretion is significantly reduced in the absence of extracellular CaCl2, which suggests that Aβ25–35-induced Ca2+ influx in platelets is required for dense granule release. We also assessed the effect of apyrase and P2Y1/P2Y12 receptor inhibitors on serotonin release (Figure 2C). Despite a trend towards reduction in the presence of MRS-2179, neither of these ADP signalling inhibitors abolished or significantly reduced dense granule release. In order to understand which signalling pathways are responsible for the Aβ25–35-dependent stimulation of dense granule secretion, we utilized different known inhibitors in [14C]serotonin secretion experiments (Figure 2D). A MAPK inhibitor (10 μM SB203580), an ERK inhibitor (10 μM PD98059), a PI3K inhibitor (20 μM LY294002), a PKC inhibitor (10 μM Ro31-8220) and a ROCK inhibitor (20 μM Y27632) were used to selectively inhibit different signalling pathways, but had no significant effect on dense granule release (Figure 2D).

25–35 induces dose-dependent α-granule and dense granule secretion

Figure 2
25–35 induces dose-dependent α-granule and dense granule secretion

(A) Flow cytometry analysis of P-selectin exposure in platelets stimulated with the indicated concentrations of Aβ25–35 (μM). Data are expressed as the percentage of positive cells, and are the means±S.E.M. of four experiments. Dense granule release is shown in (B), (C) and (D). (B) Concentration–response analysis of Aβ25–35-induced release of [14C]serotonin from washed human platelets, in the presence (1 mM CaCl2: closed bars) or in the absence (2 mM EGTA: open bars) of extracellular Ca2+. (C) Platelets were pre-treated for 10 min with 2 units/ml apyrase, 200 μM MRS-2179 or 1 μM AR-C69931MX or Hepes buffer (none) before stimulation with 20 μM Aβ25–35. Several signalling inhibitors were used in (D) to test the role of signalling pathways in the stimulation of dense granule release by Aβ25–35. A MAPK inhibitor (10 μM SB203580), an ERK inhibitor (10 μM PD98059), a PI3K inhibitor (20 μM LY294002), a PKC inhibitor (10 μM Ro31–8220) and a ROCK inhibitor (20 μM Y27632) were pre-incubated for 10 min before stimulation with 20 μM Aβ25–35. The release of serotonin in the supernatant is expressed as percentage of the total incorporated radioactivity upon subtraction of the values measured in the supernatant of resting platelets. Data are expressed as means±S.E.M. of three to five different experiments. Statistical analysis was performed using one-way ANOVA with Bonferroni post-test, n=3, *P<0.05 and **P<0.01. ns, P>0.05.

Figure 2
25–35 induces dose-dependent α-granule and dense granule secretion

(A) Flow cytometry analysis of P-selectin exposure in platelets stimulated with the indicated concentrations of Aβ25–35 (μM). Data are expressed as the percentage of positive cells, and are the means±S.E.M. of four experiments. Dense granule release is shown in (B), (C) and (D). (B) Concentration–response analysis of Aβ25–35-induced release of [14C]serotonin from washed human platelets, in the presence (1 mM CaCl2: closed bars) or in the absence (2 mM EGTA: open bars) of extracellular Ca2+. (C) Platelets were pre-treated for 10 min with 2 units/ml apyrase, 200 μM MRS-2179 or 1 μM AR-C69931MX or Hepes buffer (none) before stimulation with 20 μM Aβ25–35. Several signalling inhibitors were used in (D) to test the role of signalling pathways in the stimulation of dense granule release by Aβ25–35. A MAPK inhibitor (10 μM SB203580), an ERK inhibitor (10 μM PD98059), a PI3K inhibitor (20 μM LY294002), a PKC inhibitor (10 μM Ro31–8220) and a ROCK inhibitor (20 μM Y27632) were pre-incubated for 10 min before stimulation with 20 μM Aβ25–35. The release of serotonin in the supernatant is expressed as percentage of the total incorporated radioactivity upon subtraction of the values measured in the supernatant of resting platelets. Data are expressed as means±S.E.M. of three to five different experiments. Statistical analysis was performed using one-way ANOVA with Bonferroni post-test, n=3, *P<0.05 and **P<0.01. ns, P>0.05.

ADP release mediates the stimulation of key signalling pathways by Aβ25–35

It has been demonstrated that Aβ25–35 is able to activate intracellular signaling pathways initiating a cascade of phosphorylation of different effectors, namely the MAPKs p38MAPK and ERK1/2, PI3K and PKC [18,19]. In light of the ability of Aβ25–35 to promote granule secretion and ADP release, we investigated the possible role of ADP in platelet activation induced by Aβ25–35. Immunoblotting analysis with phospho-specific antibodies shows that Aβ25–35 induces a time-dependent phosphorylation of several key signalling proteins. These include the PI3K-regulated kinase Akt, the PKC substrate pleckstrin, ERK1/2 and p38MAPK (Figure 3A). Moreover, Aβ25–35 promotes the rapid and robust tyrosine phosphorylation of several proteins as identified using the phosphotyrosine-specific antibody 4G10 (Figure 3B). Phospho-specific bands in Figure 3(B) include a substrate with an apparent molecular mass of approximately 70 kDa, which is likely to be the non-receptor tyrosine kinase Syk. In fact, the Aβ25–35-dependent phosphorylation of Syk has been confirmed by immunoprecipitation experiments (Figure 3C). Interestingly, Aβ25–35-induced phosphorylation of these substrates is severely, yet not completely, impaired in the presence of the ADP scavenger apyrase, indicating that the positive signalling initiated by secreted ADP plays a central role in Aβ25–35-induced platelet activation. According to previous evidence [20], Aβ25–35 also induces phosphorylation of the Ca2+-regulated MLC, which, however, is only partially reduced by pre-incubation of platelets with apyrase (Figure 3D and Supplementary Figure S1 at http://www.biochemj.org/bj/462/bj4620513add.htm). This suggests that, in contrast with the molecular events described above, a significant portion of the MLC phosphorylation promoted by Aβ25–35 is ADP- and granule secretion-independent. Nonetheless, it seems likely from these data that ADP at least reinforces MLC phosphorylation in a positive feedback manner. Similar to apyrase, the inhibition of the ADP receptor P2Y12 with AR-C69931MX resulted in the strong reduction of the phosphorylation of the PI3K-regulated kinase Akt, the PKC substrate pleckstrin, ERK1/2 and p38MAPK and MLC to basal levels (i.e. no Aβ25–35) (Figure 4). Hence this confirms the crucial role of this Gi-coupled receptor in mediating the effect of secreted ADP. Phosphorylation of p38MAPK and Akt was also equally prevented by the P2Y1 receptor inhibitor MRS-2179, indicating that both receptors are involved in the mechanism of ADP potentiation of Aβ25–35-dependent phosphorylation. By contrast, P2Y1 receptor inhibition by MRS-2179 had only a partial inhibitory effect on the phosphorylation of the PKC substrate pleckstrin, ERK1/2 and MLC (Figure 4 and Supplementary Figure S2 at http://www.biochemj.org/bj/462/bj4620513add.htm), possibly suggesting a secondary role for P2Y1.

25–35-induced signalling pathway activation requires ADP release

Figure 3
25–35-induced signalling pathway activation requires ADP release

Washed platelets (5×108/ml, 0.1 ml) were treated with 10 μM Aβ25–35 for the indicated time (A, B and D) or 2 min (C). Where indicated, platelets were incubated with 2 units/ml apyrase grade VII (apyrase) or buffer (none) for 10 min at 37°C before the addition of Aβ25–35. Total proteins were separated by SDS/PAGE and protein phosphorylation was analysed by immunoblotting with the indicated antibodies: (A) anti-phospho-Akt, anti-phospho-pleckstrin (i.e. PKC activity), anti-phospho-ERK1/2, anti-phospho-p38MAPK and anti-tubulin for equal loading (P, phospho); (B) anti-phospho-Tyr (4G10); and (D) anti-phospho-MLC and anti-tubulin for equal loading. In (C), washed platelets (109/ml) treated as described above were lysed and Syk was immunoprecipitated (IP) using a specific antibody. Phosphorylation of immunoprecipitated Syk was evaluated with the anti-phospho-tyrosine antibody 4G10. The level of immunoprecipitated protein was confirmed by immunoblotting with anti-Syk antibody. Data shown are representative of three independent experiments.

Figure 3
25–35-induced signalling pathway activation requires ADP release

Washed platelets (5×108/ml, 0.1 ml) were treated with 10 μM Aβ25–35 for the indicated time (A, B and D) or 2 min (C). Where indicated, platelets were incubated with 2 units/ml apyrase grade VII (apyrase) or buffer (none) for 10 min at 37°C before the addition of Aβ25–35. Total proteins were separated by SDS/PAGE and protein phosphorylation was analysed by immunoblotting with the indicated antibodies: (A) anti-phospho-Akt, anti-phospho-pleckstrin (i.e. PKC activity), anti-phospho-ERK1/2, anti-phospho-p38MAPK and anti-tubulin for equal loading (P, phospho); (B) anti-phospho-Tyr (4G10); and (D) anti-phospho-MLC and anti-tubulin for equal loading. In (C), washed platelets (109/ml) treated as described above were lysed and Syk was immunoprecipitated (IP) using a specific antibody. Phosphorylation of immunoprecipitated Syk was evaluated with the anti-phospho-tyrosine antibody 4G10. The level of immunoprecipitated protein was confirmed by immunoblotting with anti-Syk antibody. Data shown are representative of three independent experiments.

Effect of P2Y1 and P2Y12 inhibition on Aβ25–35-induced signalling activation

Figure 4
Effect of P2Y1 and P2Y12 inhibition on Aβ25–35-induced signalling activation

Washed platelets (5×108/ml, 0.1 ml) were treated with 10 μM Aβ25–35 for 1 min. Where indicated, platelets were incubated with 2 units/ml apyrase, 200 μM MRS-2179 or 1 μM AR-C69931MX for 5 min at 37°C before the addition of Aβ25–35. Total proteins were separated by SDS/PAGE and protein phosphorylation was analysed by immunoblotting with the following antibodies: anti-phospho-Akt, anti-phospho-pleckstrin (i.e. PKC activity), anti-phospho-ERK1/2, anti-phospho-p38MAPK, anti-phospho-MLC and anti-tubulin for equal loading. Data shown are representative of three independent experiments. P, phospho.

Figure 4
Effect of P2Y1 and P2Y12 inhibition on Aβ25–35-induced signalling activation

Washed platelets (5×108/ml, 0.1 ml) were treated with 10 μM Aβ25–35 for 1 min. Where indicated, platelets were incubated with 2 units/ml apyrase, 200 μM MRS-2179 or 1 μM AR-C69931MX for 5 min at 37°C before the addition of Aβ25–35. Total proteins were separated by SDS/PAGE and protein phosphorylation was analysed by immunoblotting with the following antibodies: anti-phospho-Akt, anti-phospho-pleckstrin (i.e. PKC activity), anti-phospho-ERK1/2, anti-phospho-p38MAPK, anti-phospho-MLC and anti-tubulin for equal loading. Data shown are representative of three independent experiments. P, phospho.

Next we analysed the activation of the small GTPase Rap1b, which is an essential regulator of integrin inside-out signalling [32,33]. Using a pull-down assay we demonstrated that Aβ25–35 dose-dependently stimulates Rap1b activation (Figure 5A and Supplementary Figure S3A at http://www.biochemj.org/bj/462/bj4620513add.htm). Accumulation of Rap1b–GTP by Aβ25–35 is rapid but transient, as it peaks at 1 min (Figure 5B). The pre-treatment of platelets with 2 units/ml apyrase grade VII for 10 min at 37°C before the addition of Aβ25–35 completely abolishes the activation Rap1b (Figure 5B and Supplementary Figure S3B). The activation of Rap1b by Aβ25–35 also appears to be completely Ca2+-dependent, since it was completely inhibited by platelet pre-treatment with 30 μM BAPTA (Figure 5C and Supplementary Figure S3D). The dependence of Aβ25–35-induced Rap1b activation on ADP release and P2Y1/P2Y12 activation was further investigated using the P2Y1 inhibitor MRS-2179 and the P2Y12 inhibitor AR-C69931MX. As shown in Figure 5(D), similar to apyrase, these two inhibitors completely abolish Rap1b activation, suggesting that the activity of both receptors is necessary for the stimulation of platelets. Interestingly, the activation of Rap1b by Aβ25–35 is not modified by pre-incubation by RGDS, suggesting that in these experimental conditions Rap1b is not affected by the outside-in signalling of integrin αIIbβ3. As expected, given the pivotal role of Rap1b in stimulating integrin αIIbβ3 activation and fibrinogen binding, and in parallel to Rap1b activation, Aβ25–35 also stimulated platelet fibrinogen binding in a concentration-dependent manner (Figure 5E).

25–35 induces Rap1b activation in an ADP-dependent manner

Figure 5
25–35 induces Rap1b activation in an ADP-dependent manner

Rap1b activation (Rap1b-GTP) was evaluated by an activation state-dependent pull-down assay (GST-RalGDSRBD) followed by immunoblotting with anti-Rap1b antibody. Washed platelets were stimulated with different concentrations of Aβ25–35 (0–10 μM) for 2 min (A), for different times (0–300 s) with 10 μM Aβ25–35 with or without 10 min pre-incubation with 2 units/ml apyrase (B), pre-incubated for 2 min with 30 μM BAPTA and/or 2 units/ml apyrase for 10 min (C) or pre-incubated for 10 min with 2 units/ml apyrase, 10 μM RGDS, 200 μM MRS-2179 or 1 μM AR-C69931MX or Hepes buffer (none) before stimulation with 10 μM Aβ25–35 (D). Platelets were then lysed, and 0.5 ml of lysates was incubated with GST–RalGDSRBD. The levels of active precipitated Rap1b were tested by immunoblotting (Rap1b–GTP). Upper panel represents the immunoblot of the pull-down samples. Lower panel shows the amount of total protein (Rap1b). (E) The effect of Aβ25–35 on fibrinogen binding by platelets is analysed. Washed platelets (2×107/ml, 0.05ml) were treated with Tyrode's buffer (none) or 20 μM Aβ25–35. FITC-conjugated fibrinogen (5 μl) was added to platelet suspensions and incubated for 30 min before dilution with 0.45 ml of Hepes buffer. Fibrinogen binding was quantified by flow cytometry. Representative dot plots of fluorescent labelling (FL1-H: FBN FITC) over forward scattering (FSC) are shown in the left-hand panels, whereas the percentage of fibrinogen-positive human platelets stimulated with the indicated concentrations of Aβ25–35 are displayed in the right-hand panels. Data are expressed as means±S.E.M. of three different experiments.

Figure 5
25–35 induces Rap1b activation in an ADP-dependent manner

Rap1b activation (Rap1b-GTP) was evaluated by an activation state-dependent pull-down assay (GST-RalGDSRBD) followed by immunoblotting with anti-Rap1b antibody. Washed platelets were stimulated with different concentrations of Aβ25–35 (0–10 μM) for 2 min (A), for different times (0–300 s) with 10 μM Aβ25–35 with or without 10 min pre-incubation with 2 units/ml apyrase (B), pre-incubated for 2 min with 30 μM BAPTA and/or 2 units/ml apyrase for 10 min (C) or pre-incubated for 10 min with 2 units/ml apyrase, 10 μM RGDS, 200 μM MRS-2179 or 1 μM AR-C69931MX or Hepes buffer (none) before stimulation with 10 μM Aβ25–35 (D). Platelets were then lysed, and 0.5 ml of lysates was incubated with GST–RalGDSRBD. The levels of active precipitated Rap1b were tested by immunoblotting (Rap1b–GTP). Upper panel represents the immunoblot of the pull-down samples. Lower panel shows the amount of total protein (Rap1b). (E) The effect of Aβ25–35 on fibrinogen binding by platelets is analysed. Washed platelets (2×107/ml, 0.05ml) were treated with Tyrode's buffer (none) or 20 μM Aβ25–35. FITC-conjugated fibrinogen (5 μl) was added to platelet suspensions and incubated for 30 min before dilution with 0.45 ml of Hepes buffer. Fibrinogen binding was quantified by flow cytometry. Representative dot plots of fluorescent labelling (FL1-H: FBN FITC) over forward scattering (FSC) are shown in the left-hand panels, whereas the percentage of fibrinogen-positive human platelets stimulated with the indicated concentrations of Aβ25–35 are displayed in the right-hand panels. Data are expressed as means±S.E.M. of three different experiments.

Endogenous ADP release is necessary for Aβ25–35-induced platelet aggregation and increased thrombus formation under flow

We next analysed the ability of Aβ25–35 to stimulate platelet aggregation and investigated its mechanism of action. Platelet aggregation was induced by different concentrations of Aβ25–35 under constant stirring in a Born lumiaggregometer in the presence of 1 mM extracellular Ca2+ (Figure 6A). Aβ25–35 dose-dependently induced platelet aggregation. The effect of Aβ25–35-induced aggregation depends on Ca2+ as it is strongly reduced in the absence of extracellular Ca2+ and by pre-incubation with the intracellular Ca2+ chelator BAPTA (Figure 6B). The role of ADP secretion in Aβ25–35-induced aggregation was tested using the ADP-degrading enzyme apyrase, which strongly reduces platelet aggregation (Figure 6C), and the P2Y1/P2Y12 inhibitors MRS-2179 and AR-C69931MX (respectively), which also significantly reduced Aβ25–35-induced aggregation (Figure 6D). Notably, despite complete Ca2+ and ADP dependence of the Aβ25–35-dependent activation of the integrin αIIbβ3 activator Rap1b described above, approximately 25% of the aggregation induced by Aβ25–35 is not inhibited in the absence of Ca2+ or in the presence of ADP inhibitors. This suggests that there is a component of this response that does not depend on the activation of integrin αIIbβ3, which was directly proved using the peptide integrin inhibitor RGDS (Figure 6E). This component of the aggregatory response was also insensitive to apyrase treatment (Figure 6F), suggesting that part of the aggregation induced by Aβ25–35 is due to an integrin- and ADP-independent agglutination-like phenomenon. Interestingly, this integrin- and ADP-independent component seems responsible for the small potentiation of collagen-dependent aggregation in the presence of Aβ25–35, as concluded by experiments showing that apyrase reduces, but does not completely abolish, the effect of Aβ25–35 on collagen-dependent aggregation even in the presence of RGDS (Supplementary Figure S4 at http://www.biochemj.org/bj/462/bj4620513add.htm).

25–35-induced platelet aggregation depends on Ca2+ mobilization and ADP release

Figure 6
25–35-induced platelet aggregation depends on Ca2+ mobilization and ADP release

Human washed platelets (3×108/ml) were stimulated with different concentrations of Aβ25–35 (as indicated) in the presence of 1 mM extracellular Ca2+ (A and CF). (B) Platelets were tested in the absence of extracellular Ca2+ (none) or pre-incubated for 2 min with 30 μM BAPTA. (C) Platelets were pre-incubated for 10 min with 2 units/ml apyrase before stimulation with 10 μM Aβ25–35. (D) Platelets were treated with 200 μM MRS-2179 or 1 μM AR-C69931MX before stimulation with 10 μM Aβ25–35. (E) Platelets were incubated with 0.25 mM RGDS before stimulation with 10 μM Aβ25–35. (F) Platelets were incubated with 0.25 mM RGDS and/or 2 units/ml apyrase before stimulation with 10 μM Aβ25–35. Aggregation was monitored as increase of light transmission up to 5 min. Traces in the Figure are representative of three or more independent experiments.

Figure 6
25–35-induced platelet aggregation depends on Ca2+ mobilization and ADP release

Human washed platelets (3×108/ml) were stimulated with different concentrations of Aβ25–35 (as indicated) in the presence of 1 mM extracellular Ca2+ (A and CF). (B) Platelets were tested in the absence of extracellular Ca2+ (none) or pre-incubated for 2 min with 30 μM BAPTA. (C) Platelets were pre-incubated for 10 min with 2 units/ml apyrase before stimulation with 10 μM Aβ25–35. (D) Platelets were treated with 200 μM MRS-2179 or 1 μM AR-C69931MX before stimulation with 10 μM Aβ25–35. (E) Platelets were incubated with 0.25 mM RGDS before stimulation with 10 μM Aβ25–35. (F) Platelets were incubated with 0.25 mM RGDS and/or 2 units/ml apyrase before stimulation with 10 μM Aβ25–35. Aggregation was monitored as increase of light transmission up to 5 min. Traces in the Figure are representative of three or more independent experiments.

We have previously demonstrated that platelets are able to adhere on immobilized Aβ-peptides under static conditions and to fasten spreading over collagen [21]. In the present study we evaluated the ability of Aβ25–35 to potentiate platelet adhesion to collagen and thrombus formation under physiological flow conditions. Platelets from human whole blood were selectively labelled with calcein/AM and the formation of thrombi under arterial shear stress (1000 s−1) was assessed as described previously [29,30]. As shown in Figure 7(A), platelet adhesion and thrombus formation on collagen-coated surfaces is significantly increased by collagen co-incubation with 10 μM Aβ25–35 during coating. Surprisingly, addition of Aβ25–35 directly to blood does not increase thrombus adhesion on collagen-coated surfaces (results not shown), possibly due to increased platelet–platelet interaction rather than potentiation of adhesion to absorbed collagen. Pre-treatment of platelets with apyrase ablates the increase in thrombus formation observed in our experiments (Figure 7B), which further suggests the importance of ADP release in the potentiatory effect of Aβ25–35. Notably, we could not detect any effect of apyrase on thrombus formation on collagen-coated surfaces in the absence of Aβ25–35 (Figure 7C). This suggests that under high shear stress conditions and on collagen-coated surfaces local release of ADP and its effect on thrombus formation are limited. This also suggests that the inhibition of thrombus formation by apyrase on collagen/Aβ25–35 co-coated surface depends on the specific inhibition of an Aβ25–35-dependent event rather than an off-target inhibition of thrombus formation.

25–35 increases thrombus formation in whole blood under physiological flow conditions in an ADP-dependent manner

Figure 7
25–35 increases thrombus formation in whole blood under physiological flow conditions in an ADP-dependent manner

Human PRP was isolated from anti-coagulated blood (citrate) and incubated for 1 h with calcein/AM (5 μg/ml) at 37°C. After reconstitution of whole blood by mixing labelled PRP and the red blood cell fraction, thrombus formation was monitored under physiological shear stress conditions (1000 s−1). Coating of the microchannel was performed for 60 min at 37°C with collagen alone (1 mg/ml) (Ctrl: top microchannel in A and both channels in C) or collagen plus Aβ25–35 (20 μM) (bottom microchannel in A and both channels in B). Where indicated, apyrase (1 unit/ml) was added to the blood 10 min before the assay (B and C). Both panels show a representative picture of the thrombi and a quantification of the surface coverage after 10 min of flow (means±S.E.M., n=4). Statistical significance of the differences was measured using a Student's t test for paired samples (*P<0.05).

Figure 7
25–35 increases thrombus formation in whole blood under physiological flow conditions in an ADP-dependent manner

Human PRP was isolated from anti-coagulated blood (citrate) and incubated for 1 h with calcein/AM (5 μg/ml) at 37°C. After reconstitution of whole blood by mixing labelled PRP and the red blood cell fraction, thrombus formation was monitored under physiological shear stress conditions (1000 s−1). Coating of the microchannel was performed for 60 min at 37°C with collagen alone (1 mg/ml) (Ctrl: top microchannel in A and both channels in C) or collagen plus Aβ25–35 (20 μM) (bottom microchannel in A and both channels in B). Where indicated, apyrase (1 unit/ml) was added to the blood 10 min before the assay (B and C). Both panels show a representative picture of the thrombi and a quantification of the surface coverage after 10 min of flow (means±S.E.M., n=4). Statistical significance of the differences was measured using a Student's t test for paired samples (*P<0.05).

DISCUSSION

The ability of Aβ-peptides to promote activation and adhesion of platelets has been previously reported by our group and others [1821,34], and is likely to significantly contribute to the development and progression of cerebrovascular diseases associated with AD. In the present study, we investigated the molecular mechanism underlying the activation of platelets by Aβ-peptides, and we demonstrated for the first time the crucial and hierarchical role of intracellular Ca2+ increase and ADP secretion. We propose that Ca2+-mediated dense granule release and secretion of ADP is an early event in platelet stimulation by Aβ25–35, and that secreted ADP is the major player in the stimulation and propagation of platelet activation. We demonstrated that pre-incubation of platelets with the ADP-scavenging enzyme apyrase strongly reduces Aβ25–35-induced platelet aggregation and potentiation of thrombus formation. Activation of the small GTPase Rap1b, and consequentially integrin αIIbβ3 stimulation, are also dependent on ADP. We have shown that several intracellular signalling pathways (namely p38MAPK, ERK, PI3K and PKC) are activated by Aβ25–35, although they do not play a role in the stimulation of dense granule and ADP release. Moreover, despite a marginal ADP-independent component, Aβ25–35-induced phosphorylation and activation of the signalling proteins Syk, PKC, PI3K and p38MAPK and ERK1/2 heavily depends on ADP, since this is significantly reduced in the presence of apyrase and P2Y receptor inhibitors. The results of the present study are in substantial agreement with our previous studies showing that in platelets adhering to immobilized Aβ-peptides the release of ADP reinforces the activation of intracellular signalling pathways [21].

The results of the present study clearly show that Aβ25–35 induces a significant increase in intracellular Ca2+ that is necessary for the stimulation of Aβ25–35-induced α-granule and dense granule release, and is critical for platelet aggregation. This observation is consistent with previous hypotheses that Aβ-peptides can generate pores in the plasma membrane of cells leading to Ca2+ influx from the extracellular space [22,23,35]. However, it remains unclear at the present time whether Aβ25–35 generates pores in the plasma membrane of human platelets to allow Ca2+ influx or whether it regulates pre-existing Ca2+ channels. Hence, the molecular mechanisms linking Aβ-peptides to platelet intracellular Ca2+ increase remain elusive and will require further investigation.

We documented the activation of PKC in Aβ25–35-stimulated platelets, as revealed by the phosphorylation analysis of the intracellular protein pleckstrin [36]. The activation of PKC {which is usually downstream of PLC (phospholipase C) activation, Ins(1,4,5)P3 generation and release of Ca2+ from intracellular stores [37]} is not an early event in platelet activation by Aβ-peptide. In fact, unlike what we observed for the intracellular Ca2+ increase, the complete inhibition of pleckstrin phosphorylation by the ADP scavenger apyrase demonstrates that PLC/PKC activation is secondary to granule release. We have also shown that the inhibition of PKC by Ro31-8220 does not affect dense granule release by Aβ25–35. These observations are in partial disagreement with a previous study indicating that inhibition of PKC by Ro31-8220 prevents Aβ25–35-induced Ca2+ increase [19]. Although it is difficult to understand the reason for this discrepancy, we would like to highlight that the inhibition of the Ca2+ spike shown by those authors is only partial, which does not exclude the possibility of a double source for the intracellular Ca2+ increase in response to Aβ25–35 (i.e. partially from membrane permeabilization and partially from intracellular store release). Moreover, the limited specificity of Ro31-8220 could be responsible for some misinterpretation of the data in the above study [38].

Recently, an important early role for the RhoA-activated protein kinase ROCK in platelet activation by Aβ25–35 has been proposed [20]. Although the ROCK inhibitor Y27632 did not inhibit the Ca2+ wave or the dense granule release induced by Aβ-peptides, the results in the present study are not in disagreement with that study. In fact, although we prove that ROCK is not responsible for the initial increase in intracellular Ca2+ and release of ADP, our data do not exclude that ROCK is indeed stimulated by Aβ-peptides and plays an important role in the resulting platelet activation. This is in agreement with our data showing that MLC (a known substrate for ROCK) is phosphorylated in response to platelet incubation with Aβ25–35.

One of the strengths of the results of the present study is in the identification of a strong intracellular Ca2+-dependent stimulation of granule secretion in the presence of Aβ-peptides. Interestingly, upon stimulation with Aβ-peptides, MLC is rapidly phosphorylated, which is likely to result in the reorganization of actomyosin contractility and the release of α-granules and dense granules [39]. The inhibition of ADP in our cell signalling experiments severely impaired the phosphorylation of all of the intracellular targets analysed, yet it only marginally reduced the phosphorylation of MLC, suggesting that this cellular event occurs upstream of granule secretion, and may be directly driven by intracellular Ca2+ increase. On the other hand, the activation of the other protein kinases investigated in the present study and their signalling is likely to be a consequence of granule and ADP release.

One limitation of the present and most in vitro studies on the effect of Aβ-peptides on platelets is the chosen concentration of the peptides [1821,34]. In all cases, the concentration chosen for platelet studies is within the 1–20 μM range, whereas the concentration in human tissues appears significantly lower, ranging between 0.2 and 40 nM [40,41]. This is often the case for platelet studies performed in vitro. For example, ADP studies are performed with concentrations ranging between 10 and 50 μM, whereas ADP in the plasma is at submicromolar concentrations [42]. Besides a possible reduced efficacy of agonists and other biomolecules in vitro, it is important to bear in mind that systemic or macroscopic tissue concentration measurements are a gross underestimation of the local concentrations of physiological molecules in their microenvironment. Moreover, the utilization of higher concentrations for studies in vitro is likely to facilitate the clarification of molecular mechanisms otherwise very difficult to tease out with low/physiological concentrations.

Taken together, our data suggest the centrality of ADP in the stimulation of platelet activation, aggregation and thrombus formation by Aβ-peptides. We propose a signalling model for Aβ-peptide stimulation of platelet responses, in which Ca2+ influx into the platelet leads to dense granule release and ADP secretion. Consequently, ADP-dependent activation of signalling through several intracellular protein kinase pathways and activation of Rap1b plays a prominent role in the stimulation of platelet aggregation and thrombus formation. As shown previously [43], the activation of Rap1b leads to integrin inside-out signalling and stimulates fibrinogen binding at the base of key physiological phenomena such as platelet aggregation and thrombus formation. To our knowledge, our novel observations have the potential to clarify the pathological role of Aβ-peptides in cerebrovascular disease and indicate that platelet ADP receptors are prospective novel targets for the treatment of vascular complications of AD.

Abbreviations

     
  • amyloid β

  •  
  • AD

    Alzheimer’s disease

  •  
  • AM

    acetoxymethyl ester

  •  
  • APP

    amyloid precursor protein

  •  
  • BAPTA

    1,2-bis-(o-aminophenoxy)ethane-N,N,N′,N′-tetra-acetic acid

  •  
  • ERK

    extracellular-signal-regulated kinase

  •  
  • MAPK

    mitogen-activated protein kinase

  •  
  • MLC

    myosin light chain

  •  
  • PI3K

    phosphoinositide 3-kinase

  •  
  • PKC

    protein kinase C

  •  
  • PLC

    phospholipase C

  •  
  • PRP

    platelet-rich plasma

  •  
  • RGDS

    Arg-Gly-Asp-Ser

  •  
  • ROCK

    Rho-associated kinase

  •  
  • sAPP

    soluble APP

AUTHOR CONTRIBUTION

Ilaria Canobbio performed most of the experiments. Gianni Guidetti and Daria Manganaro performed analysis of secretion. Barbara Oliviero performed flow cytometry experiments. Dina Vara performed thrombus formation assays. Ilaria Canobbio, Mauro Torti and Giordano Pula designed the experiments, analysed the data and wrote the paper.

We thank Caterina Visconte (Department of Biology and Biotechnology, University of Pavia, Pavia, Italy) and Dr Laura Di Pasqua (Department of Internal Medicine and Therapeutics, University of Pavia, Pavia, Italy) for technical support.

FUNDING

This work was supported by ARUK (Alzheimer's Research UK) [grant number PPG2013B-8 (to G.P.)].

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Author notes

2

Joint senior authors.

Supplementary data